Training Curriculum of Laboratory
Standards for Primary Health Care Clinics
in Iraq
June 2013
List of Contents
List of contents
List of abbreviations
Introduction
Part One: Trainers guide
How to use the manual
Structure of the training course
Approaches to training and learning
Part Two: Training modules
Module1: Introduction, Setting up a primary health care laboratory, and safety in the
laboratory
Session1: Introduction and Setting up a primary health care laboratory
Session 2: Safety in the laboratory
Module 2: Hematology
Session 1: Collection and handling of blood and automated blood count techniques
Session 2: Manual techniques
Module 3: Serology
Session 1: RPR Test, TPHA Test, Widel Test, and Rose Bengal Test
Session 2: CRP Test, ASO latex slide agglutination Test, RF, Toxoplasmosis Test,
Infectious mononucleosis Test, Rota Virus Latex Test, and Pregnant Test
Module 4: Bacteriology
Session 1: General urine examination and general stool examination
Session 2: Urine culture, throat swab, and nasal swab
Session 3: Smart tests, preparation of media and staining smears
Module 5: Parasitology and biochemistry
Session 1: Parasitology
Session 2: Biochemistry
Annex 1
References
1
2
5
6
7
8
9
39
40
41
45
60
61
88
119
120
137
156
157
186
211
233
234
243
274
284
Page | 2
Acronyms
AAFB
AIDS
ALP
ALT
AST
BHSP
CRP
CSF
CV
DNA
EDTA
ELISA
ESR
GIT
Hb
HCG
Hct
Hi
HiCN
HIV
IDDM
Ig
IHA
IU
KIA
LDH
LPS
MAC
MCH
MCV
MIU
MoH
MPV
MPXI
MRSA
NIDDM
PCV
PDW
PHC
PPE
R
Acid and Alcohol Fast Bacilli
Acquired Immune Deficiency Syndrome
Alkaline Phosphatase
Alanine Aminotransferase
Aspartate Aminotransferase
Basic Health Services Package
C- Reactive Protein
Cerebro-Spinal Fluid
Coefficient of Variation
Deoxyribonucleic Acid
Ethylenediaminetetraacetic Acid
Enzyme-Linked Immunosorbent Assay
Erythrocyte Sedimentation Rate
Gastro Intestinal Tract
Hemoglobin
Human Chorionic Gonadotropin
Hematocrit
Meth hemoglobin
Hemiglobincyanide
Human Immune Virus
Insulin Dependent Diabetes Miletus
Immunoglobulin
Indirect Haemogglutination
International Unit
Klinger Iron Agar
Low Density Lipids
Lipopolysaccharides
MacConky
Mean Corpuscular Hemoglobin
Mean Corpuscular Volume
Motility-Indole-Urase
Ministry of Health
Mean Platelet Volume
Mean Peroxidase Activity Index
Methicillin Resistant Staph Aurous
Non-Insulin Dependent Diabetes Mellitus
Packed Cell Volume
Platelet Distribution Width
Primary Health Care
Personal Protective Equipment
Reactive
Page | 3
RBCs
RDW
RF
RNA
RPM
RPR
SD
SHb
SOPs
SPA
SR
TCBS
TNCC
TPHA
TSA
URC
USAID
UTI
VCA
VDRL
WBC
XLD
Red Blood Cells
Red cell Distribution Width
Rheumatoid Factor
Ribonucleic Acid
Round Per Minute
Rapid Plasma Reagin
Standards Deviation
Sulphemoglobin
Standard Operating Procedures
Suprapubic Aspirate
Strong Reactive
Thiosulfate Citrate Bile Salts
Total Nucleated Cell Count
Treponema Pallidum Particle Agglutination
Tryptic Soya Agar
University Research Company
United States Agency of International Development
Urinary Tract Infection
Viral Structural Antigens
Venereal Disease Research Laboratory
White Blood Cells
Xylose Lysine Desoxycholate
Page | 4
Introduction
The Iraqi Ministry of Health (MoH) is undergoing health reform and moving forward with
reorganizing and stabilizing its health services. Ensuring access to routine, high quality, and
equitable healthcare has emerged as a critical need, and the Government of Iraq (GoI) is
responding with vigor and commitment to improving the quality of primary health care (PHC)
services. To assist with these efforts, USAID awarded University Research Co. LLC (URC)
the four-year Primary Health Care Project in Iraq (USAID/PHCPI) to support the MoH in
achieving its goal of improving the quality of care for all Iraqis. The project aims to strengthen
health sector capacity to provide essential PHC services and expand availability of such
services throughout the country, with increasing coverage for vulnerable and at-risk
populations, such as children under five, women, and internally displaced persons.
The "Laboratory Standard Operating Procedures," often called the "Methods," discusses how
the experiment occurred. Documenting the procedures of the laboratory experiment is
important not only so that others can repeat the results but also so that you can replicate the
work later, if the need arises. Historically, laboratory procedures have been written as firstperson narratives as opposed to second-person sets of instructions.
Achieving a proper depth in laboratory procedures is challenging. In general, adequate
information should be given in order to let the staff able to do the procedure. For that reason,
details that will affect the outcome are included.
The Basic Health Services Package for Iraq was considered in developing this guideline as all
the SOPs for the investigations are listed in the BHSP.
Page | 5
Part 0ne
Trainer’s Guide
Page | 6
Part I: Trainer’s Guide
This training curriculum is a guide to assist trainers in improving health care by training health
professionals by applying laboratory standards in primary health care centers
Materials in this document are designed for training service providers who work at a variety
of health facilities in Iraq. The modules can be used to train health professionals, physicians
and other health workers in group training or, with adaptation, as a basis of individualized or
self-directed learning.
Trainers implementing this course should be thoroughly familiar with the guideline. The
trainers need to have a positive attitude about the participants and their training work.
Training may be implemented either off-site or on-site. In off-site training, a group of
participants come together from several health facilities and then return to apply what has been
learned. Off-site training may be the most appropriate way to reach individuals from many
small sites. On-site training refers to training held in a health facility team where the
participants work. Both types of training can be very effective. When training is conducted offsite, it may be more difficult to observe actual clinical settings. On the other hand, when
training takes place on-site, there may be interruptions due to participants being called away
for other responsibilities.
How to Use the Manual
This manual is designed as a working instrument for trainers and facilitators. The module
schedule contains a condensed summary of the contents organized in units and is meant as a
check list for the facilitator/s before and during the course. The time indicated for each unit is
an average time span based on experience, and can vary according to the composition and
dynamics of each respective group.
The manual is divided into two parts. The first part is an introduction to the training course
giving an overview over the rationale, objectives, and target groups for the course. It includes
the present section on recommendations on how to use the manual, introducing the structure,
training methods and course schedule. It also contains information on how to organize a
workshop / training course and concludes with some recommendations on the limitations of
the document and how to deal with them.
The second part presents the actual training contents, methods, didactic materials and
additional literature recommended for each content area, organized/compiled in the different
modules of the program. Every training course starts with the introduction of participants and
team presenting the course objectives, contents, methods and program and allowing
participants to express their expectations and fears.
The course content is presented according to three broad content areas (modules), subdivided
into different sessions:
Page | 7
Overall learning objectives: states the objectives to be achieved at the end of the module in
terms of knowledge, skills and competence.
Schedule: gives an overview over the time span, methods, materials and recommended content
for each session / topic and states the specific objectives of each session.
Sessions: are subdivisions/sessions of the module that follow a logical flow to develop the
content of the module.
Specific objectives of the sessions: relate to the content and the expected level of competence
to be achieved and can also be used as basis for the development of exam questions.
Background informationfor the facilitator: includes background information important for
the facilitator to develop the content of the module, necessary and recommended definitions,
concepts, theory and its applications.
Exercises: describe practical applications of the theory and are meant to facilitate the learning
process through experiential approaches: role plays, games, etc. (see list of exercises).
Handouts: are the essential documentation for the participants about the content of the session
/ module stating the objectives, listing the key words, developing the concept / theory of the
content, and giving recommendations for further reading.
References: additionally recommended literature, articles and books, which are related to the
content of the module.
Structure of the Training Course
The training course has been planned as a three days course. However, it is also possible to
shorten the course due to limited time and / or to select modules according to learning
objectives and needs. As well the time can be expanded in order to deal more in depth with the
content and allow for more exercises, practical, field work.
The time frame of the training course consists of six working hours per day. These hours are
divided into two morning and two afternoon sessions. Each session normally has duration of 2
hours. The number of course trainers/ facilitators can range from one to two per course
according to the requirements. Also, for special topics, external resource persons should be
asked to lecture and work with the group in their respective areas of expertise. The trainee facilitator ratio should be 15 to one, a ratio of 20 or 25 to one still being acceptable. The total
number of participants should not exceed 25.
The course structure and training methods not only allow for the development of knowledge,
skills, competence and change of attitudes of the participants. The course concept is also
Page | 8
designed to be put into practice by participants after the training during their work or by
organizing their own training courses.
Approaches to Training and Learning
The training course outlined in this document is based on adult learning principles,
competency-based training and performance improvement. Selected elements of the
strategies that guided the development of this material and should guide its implementation
and use are listed below.
How people learn best
People learn best when the following conditions are met:








Participants are motivated and not anxious, know what is expected of them and treated
with respect
Information and skills are interesting, exciting, meaningful, and build on what
participants already know, encourage problem-solving and reasoning
Experiences are organized, logical, practical, include a variety of methods, and protocols
and procedures are available
New learning experiences are relevant to work and training needs of participants, and are
applied immediately
Training involves every participant in active practice and participants share responsibility
for learning
Training is a team activity, including trainers and co-trainers, providing participants with
a variety of experiences and limiting trainer’s biases
The trainer acts as a facilitator of the learning process rather than a teacher who “spoon
feeds” the learner
The role and responsibilities of the trainers/facilitators and those of the
participants/learners are clearly defined with:
o The facilitators responsible for providing the learners with the necessary opportunities
to acquire the knowledge and skills necessary to perform the tasks for which they are
being trained
o The facilitators responsible for providing the learners with the necessary opportunities
to be exposed to the attitudes necessary to implement the acquired skills in a
systematic manner and initiate the process of internalizing these attitudes
o The learner remains responsible for her/his learning
The transactional relationships between the learners and the facilitators are at the level of
adult to adult characterized by mutual respect and support

Trainers are knowledgeable and competent in the subject matter and skills, use a variety
of training methods, pay attention to individual participants’ concerns, and provide
motivation through feedback and reinforcement
Page | 9

Participants must be selected according to specific criteria, such as the relevance of the
training content to the job expectations/tasks

Participants must have the necessary prerequisite level to enable them to benefit from the
learning experience

Feedback is immediate and focused on behavior that the participants can control

Assessment of learning and skills is based on objectives that the participants understand
Knowledge, skills and attitudes
This course aims to improve health care by changing health workers’ knowledge, skills and
attitudes.
 Knowledge includes the facts that the participants need to know to perform their
jobs.
Tips on increasing knowledge through training





Start with what the participants already know or have experienced
Use a variety of educational resources, including participatory activities that require
participants to use their knowledge
Use learning aids
Review and summarize often
Assess knowledge to verify learning
 Skills include the specific tasks that participants need to be able to perform.
Tips on increasing skills through training






Describe the skill
Provide protocols and procedures
Demonstrate the skill
Have participants demonstrate the skill
Verify that each skill is practiced correctly
Assess skill by observation using a checklist
 Attitudes affect behaviors, such as whether learned skills are applied and
interactions with clients.
Page | 10
Tips on changing attitudes and behavior through training






Provide information and examples
Include direct experience
Invite discussion of values, concerns and experience
Use role plays and brainstorming
Model positive attitudes
Assess changes in attitude by observing behavior
Methods
The training will use a participatory and “hands on” approach where the role of the trainers is
to facilitate learning by the participants. The responsibility for learning remains with the
participants.
Participants learn more and stay engaged in learning activities when they play an active role
in their learning and a variety of training methods are used. The following methods are
recommended in the curriculum/modules.
Selected Training Methods
Brainstorming
Case study
Clinical session
Demonstration
Discussion
Field visits
Plenary group exercises
Group assignments
Individual assignments
Individual exercises
Interview
Lecture-Discussion
Mini-lecture
Observations
Pairs exercises
Presentation
Questions and answers
Research
Return demonstration
Role play
Self-directed activities
Small group discussion
Simulation
Small group exercises
Summary
Survey
Team building exercises
Page | 11
In each module or session
This document contains an outline of a training plan for each of the key areas of content.
Each module contains the following sections:

Front page with a module number, module objectives, module content by session and an
estimated duration for the module.

Session plans covering the various content areas.
Each session contains the following sections:

Trainer Preparation: This section lists the specific preparations that trainers should
make for the session. Preparations for every session include:
 Making sure the room is properly arranged
 Ensuring that markers and flip chart or a writing board with chalk or markers are
available
 Reviewing the training plan
 Reviewing steps for the methods used in the training session
 Ensuring that the resources needed to facilitate the learning process are available
including copying materials that participants need

Methods and Activities: This section lists the methods and activities that are used in the
module. General instructions for methods that are frequently used are included in this
introductory material. Instructions for participatory activities are included in the training
plan.

Resources: The relevant reference materials/handouts and other resources needed are
listed here.

Evaluation/assessment: Evaluation methods for the knowledge or skills included are
listed. Questionnaires and skills checklists are included where needed.

Estimated Time: The time that each session/module will require depends upon the
particular group of participants, the amount of time available and other constraints. The
module gives a general time range to allow for flexible scheduling.

Training Plan: This section gives the specific learning objectives or purpose of a
session, the key ”must know” content, and the appropriate training methods and
activities for each objective. All modules include one or more activities that give
participants structured, participatory practice with the content of the module.

Handouts: When specific activities require handouts, these are included after the training
plan and should be copied before the session in which they will be used
Page | 12

Skills Checklists: Each session that includes skills objectives includes a skills
assessment checklist. The checklist is used by the trainer to evaluate the participant’s skill
based on observation of the specific steps included in the skill. The skills checklists are
also used by each participant to assess their performance and take charge of their own
learning. They can also be used by other participants for peer assessment. It is
recommended that these checklists not only be used during training to assess the
acquisition of skills, but also for post training evaluation and supervision.
Note: There are various possible formats for modules and sessions. Provided the necessary
information is included for the trainer to use, the selection of format will depend on how
comfortable the trainers are in using it.
Methods:Instructions for methods used frequently in this training course are included here. Activities
for specific methods are included with the sessions where they are used.
Mini-lecture
Trainer makes a short (5 to 15minutes) presentation using the materials available. Minilectures are used to provide information and knowledge. They insure that all participants
have an adequate level of information and insure standardization and uniformity of this
information. Mini-lectures should be kept short and should be followed by questions and
answers for clarification to enable participants to better understand the content of the
session/module and clarify issues, and questions and answers for evaluation to ensure
comprehension.
Questions and Answers (Q&A)
Questions and answers sessions are used to recall information or elicit participants’
knowledge (in introductory sessions in order to assess training needs), for clarification (to
ensure that participants understand information/content), presentation of information (to elicit
information that participants may already know) and evaluation (to assess acquisition of
knowledge and fill gaps in participants’ knowledge).
Steps for Questions and Answers for clarification
1.
2.
3.
4.
Trainer asks participants if they have questions
If a participant has a question, trainer asks another participant to answer
If the participant’s answer is correct and complete, trainer reinforces
If the participant’s answer is incorrect and/or incomplete, trainer may ask questions
that lead the participant to a more correct answer or ask another participant to answer
5. If the answer is still incorrect and/or incomplete after two or three trials, trainer
corrects and/or completes and informs the participants where to find the information
6. If there are no questions, trainer asks questions to verify knowledge and follows the
same steps (3, 4, 5)
Page | 13
Steps for Questions and Answers to elicit information from participant (s)
1. Trainer asks participants questions
2. If a participant’s answer is correct and complete, trainer reinforces
3. If the participant’s answer is incorrect and/or incomplete, trainer may ask questions
that lead the participant to a more correct answer or ask another participant to answer
4. If the answer is still incorrect and/or incomplete after two or three trials, trainer
corrects and/or completes and informs the participants where to find the information
Brainstorming
Brainstorming is an excellent way to find out what participants already know and gaps in
their knowledge. Brainstorming brings participants experience into the classroom and lets the
participants know that their experience is valuable.
Brainstorming is also a very effective way for problem solving.
A brainstorming session should always end with a summary.
Steps for brainstorming
1. Trainer asks an open-ended question
2. Participants shout out their answers or ideas:
 Until no more ideas are generated, or at least every participant has a chance to
 contribute or time allocated has run out
 No ideas are discarded criticized or analyzed, but clarifying questions can be
 asked
3. Trainer records ideas on newsprint or in another format where all can see them
How to Use the Manual
This manual is designed as a working instrument for trainers and facilitators. The module
schedule contains a condensed summary of the contents organized in units and is meant as a
checklist for the facilitator/s before and during the course. The time indicated for each unit is
an average time span based on experience, and can vary according to the composition and
dynamics of each respective group.
The manual is divided into two parts. The first part is an introduction to the training course
giving an overview over the rationale, objectives, and target groups for the course. It includes
the present section on recommendations on how to use the manual, introducing the structure,
training methods and course schedule. It also contains information on how to organize a
workshop / training course and concludes with some recommendations on the limitations of
the document and how to deal with them.
Page | 14
The second part presents the actual training contents, methods, didactic materials and
additional literature recommended for each content area, organized/compiled in the different
modules of the program. Every training course starts with the introduction of participants and
team presenting the course objectives, contents, methods and program and allowing
participants to express their expectations and fears.
The course content is presented according to three broad content areas (modules), subdivided
into different sessions:
Overall learning objectives: states the objectives to be achieved at the end of the module in
terms of knowledge, skills and competence.
Schedule: gives an overview over the time span, methods, materials and recommended content
for each session / topic and states the specific objectives of each session.
Sessions: are subdivisions/sessions of the module that follow a logical flow to develop the
content of the module.
Specific objectives of the sessions: relate to the content and the expected level of competence
to be achieved and can also be used as basis for the development of exam questions.
Background information for the facilitator: includes background information important for
the facilitator to develop the content of the module, necessary and recommended definitions,
concepts, theory and its applications.
Exercises: describe practical applications of the theory and are meant to facilitate the learning
process through experiential approaches – role-plays, games, etc.… (See list of exercises).
Handouts: are the essential documentation for the participants about the content of the session
/ module stating the objectives, listing the key words, developing the concept / theory of the
content, and giving recommendations for further reading.
References: additionally recommended literature, articles and books, which are related to the
content of the module.
Structure of the Training Course
The training course has been planned as a three days course. It is possible to shorten the course
due to limited time and / or to select modules according to learning objectives and needs. The
time can also be expanded in order to deal more in depth with the content and allow for more
exercises, practical, fieldwork.
Page | 15
The time frame of the training course consists of six working hours per day. These hours are
divided into two mornings and two afternoon sessions. Each session normally has duration of
two hours. The number of course trainers/ facilitators can range from one to two per course
according to the requirements. Also, for special topics, external resource persons should be
asked to lecture and work with the group in their respective areas of expertise. The trainee facilitator ratio should be 15 to one, a ratio of 20 or 25 to one still being acceptable. The total
number of participants should not exceed 25.
The course structure and training methods not only allow for the development of knowledge,
skills, competence and change of attitudes of the participants. The course concept is designed
for participants to put into practice after the training during their work or by organizing their
own training courses.
Approaches to Training and Learning
The training course outlined in this document is based on adult learning principles,
competency-based training and performance improvement. Selected elements of the strategies
that guided the development of this material and should guide its implementation and use are
listed below.
How People Learn Best
People learn best when the following conditions are met:







Participants are motivated and not anxious, know what is expected of them and treated with
respect
Information and skills are interesting, exciting, meaningful, and build on what participants
already know, encourage problem-solving and reasoning
Experiences are organized, logical, practical, include a variety of methods, and protocols
and procedures are available
New learning experiences are relevant to work and training needs of participants, and are
applied immediately
Training involves every participant in active practice and participants share responsibility
for learning
Training is a team activity, including trainers and co-trainers, providing participants with
a variety of experiences and limiting trainer’s biases
The trainer acts as a facilitator of the learning process rather than a teacher who “spoon
feeds” the learner
Page | 16

The role and responsibilities of the trainers/facilitators and those of the participants/learners
are clearly defined with:
o The facilitators responsible for providing the learners with the necessary opportunities
to acquire the knowledge and skills necessary to perform the tasks for which they are
being trained
o The facilitators responsible for providing the learners with the necessary opportunities
to be exposed to the attitudes necessary to implement the acquired skills in a systematic
manner and initiate the process of internalizing these attitudes
o The learner remains responsible for her/his learning
The transactional relationships between the learners and the facilitators are at the level of adult
to adult characterized by mutual respect and support.





Trainers are knowledgeable and competent in the subject matter and skills, use a variety of
training methods, pay attention to individual participants’ concerns, and provide
motivation through feedback and reinforcement
Participants must be selected according to specific criteria, such as the relevance of the
training content to the job expectations/tasks
Participants must have the necessary prerequisite level to enable them to benefit from the
learning experience
Feedback is immediate and focused on behavior that the participants can control
Assessment of learning and skills is based on objectives that the participants understand
Knowledge, Skills and Attitudes
This course aims to improve health care by changing health workers’ knowledge, skills and
attitudes.
 Knowledge includes the facts that the participants need to know to perform their
jobs.
Tips on increasing knowledge through training





Start with what the participants already know or have experienced
Use a variety of educational resources, including participatory activities that require
participants to use their knowledge
Use learning aids
Review and summarize often
Assess knowledge to verify learning
Page | 17
 Skills include the specific tasks that participants need to be able to perform.
Tips on increasing skills through training






Describe the skill
Provide protocols and procedures
Demonstrate the skill
Have participants demonstrate the skill
Verify that each skill is practiced correctly
Assess skill by observation using a checklist
 Attitudes affect behaviors, such as whether learned skills are applied and
interactions with clients.
Tips on changing attitudes and behavior through training






Provide information and examples
Include direct experience
Invite discussion of values, concerns and experience
Use role plays and brainstorming
Model positive attitudes
Assess changes in attitude by observing behavior
Methods
The training will use a participatory and “hands on” approach where the role of the trainers is
to facilitate learning by the participants. The responsibility for learning remains with the
participants.
Participants learn more and stay engaged in learning activities when they play an active role in
their learning and a variety of training methods are used. The following methods are
recommended in the curriculum/modules.
Page | 18
Selected Training Methods
Brainstorming
Case study
Clinical session
Demonstration
Discussion
Field visits
Plenary
group
exercises
Group assignments
Individual assignments
Individual exercises
Interview
Lecture-Discussion
Mini-lecture
Observations
Pairs exercises
Presentation
Questions and answers
Research
Return demonstration
Role play
Self-directed activities
Small group discussion
Simulation
Small group exercises
Summary
Survey
Team building exercises
In each Module or Session
This document contains an outline of a training plan for each of the key areas of content.
Each module contains the following sections:


Front page with a module number, module objectives, module content by session
and an estimated duration for the module.
Session plans covering the various content areas.
Each session contains the following sections:

Trainer Preparation: This section lists the specific preparations that trainers
should make for the session. Preparations for every session include:
 Making sure the room is properly arranged
 Ensuring that markers and flip chart or a writing board with chalk or markers
are available
 Reviewing the training plan
 Reviewing steps for the methods used in the training session
 Ensuring that the resources needed to facilitate the learning process are
available including copying materials that participants need

Methods and Activities: This section lists the methods and activities that are used
in the module. General instructions for methods that are frequently used are
included in this introductory material. Instructions for participatory activities are
included in the training plan.

Resources: The relevant reference materials/handouts and other resources needed
are listed here.
0|Page

Evaluation/assessment: Evaluation methods for the knowledge or skills included
are listed. Questionnaires and skills checklists are included where needed.

Estimated Time: The time that each session/module will require depends upon the
particular group of participants, the amount of time available and other constraints.
The module gives a general time range to allow for flexible scheduling.

Training Plan: This section gives the specific learning objectives or purpose of a
session, the key “must know” content, and the appropriate training methods and
activities for each objective. All modules include one or more activities that give
participants structured, participatory practice with the content of the module.

Handouts: When specific activities require handouts, these are included after the
training plan and should be copied before the session in which they will be used.

Skills Checklists: Each session that includes skills objectives includes a skills
assessment checklist. The checklist is used by the trainer to evaluate the
participant’s skill based on observation of the specific steps included in the skill.
Each participant is to use the skills checklists to assess their performance and to
take charge of their own learning. Other participants can also use the skill checklists
for a peer assessment. It is recommended that these checklists should be used for
both during training to assess the acquisition of skills and also for post-training
evaluation and supervision.
Note: There are various possible formats for modules and sessions. The format selected
will depend on how the necessary information provided and the how comfortable the
trainers are in using it.
Methods:
Instructions for methods used frequently in this training course are included here.
Activities for specific methods are included with the sessions where they are used.
Mini-lecture
Trainer makes a short (5 to 15 minutes) presentation using the materials available. Minilectures are used to provide information and knowledge. They insure that all
participants have an adequate level of information and insure standardization and
uniformity of this information. Mini-lectures should be kept short and should be
followed by questions and answers for clarification to enable participants to better
understand the content of the session/module and clarify issues, and questions and
answers for evaluation to ensure comprehension.
1|Page
Questions and Answers (Q&A)
Q&A sessions are used to recall information or elicit participants’ knowledge (in
introductory sessions in order to assess training needs), for clarification (to ensure that
participants understand information/content), presentation of information (to elicit
information that participants may already know) and evaluation (to assess acquisition
of knowledge and fill gaps in participants’ knowledge).
Steps for Questions and Answers for Clarification
1.
2.
3.
4.
Trainer asks participants if they have questions
If a participant has a question, trainer asks another participant to answer
If the participant’s answer is correct and complete, trainer reinforces
If the participant’s answer is incorrect and/or incomplete, trainer may ask
questions that lead the participant to a more correct answer or ask another
participant to answer
5. If the answer is still incorrect and/or incomplete after two or three trials, trainer
corrects and/or completes and informs the participants where to find the
information
6. If there are no questions, trainer asks questions to verify knowledge and follows
the same steps (3, 4, 5)
Steps for Questions and Answers to Elicit Information from Participant(s)
1. Trainer asks participants questions
2. If a participant’s answer is correct and complete, trainer reinforces
3. If the participant’s answer is incorrect and/or incomplete, trainer may ask
questions that lead the participant to a more correct answer or ask another
participant to answer
4. If the answer is still incorrect and/or incomplete after two or three trials, trainer
corrects and/or completes and informs the participants where to find the
information
Brainstorming
Brainstorming is an excellent way to find out what participants already know and the
gaps in their knowledge. Brainstorming brings participants experience into the
classroom and lets the participants know that their experience is valuable.
Brainstorming is also a very effective way for problem solving.
A brainstorming session should always end with a summary.
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Steps for Brainstorming
1. Trainer asks an open-ended question
2. Participants shout out their answers or ideas:
 Until no more ideas are generated, or at least every participant has a chance
to contribute or the time allocated has run out
 No ideas are discarded, criticized or analyzed, but clarifying questions can
be asked
3. Trainer records ideas on newsprint or in another format where all can see them
4. Trainer leads a discussion of each of the ideas generated
5. Trainer clearly marks ideas that are agreed upon
6. Trainer summarizes or asks participants to summarize points of agreement
7. Trainer moves to the next question only after finishing discussion of previous
question
8. Ideas generated in brainstorming can be used for summarizing, as input to group
exercises, and to relate content to participant experience
Case Study
A case study is method of training. Data and information about a case, preferably a real
one or based on one, is presented to the participants for review and analysis. It includes
specific questions to be answered. Case studies are an effective way to allow
participants to practice using information to solve problem, the highest level of
knowledge objective. They are also effective in providing participants opportunities to
explore their attitudes and confront/compare them with other participants and trainers’
attitudes. Moreover, case studies allow for the identification of gaps in knowledge.
Participants, individually or in small groups are asked to study the case and prepare
responses to the questions. The responses are then processed. During the processing the
trainer must encourage and ensure that all participants get a chance to provide inputs.
Processing can be done using questions and answers and/or discussion.
The questions must be answered in an orderly manner in the sense that each question
must be answered fully and the inputs summarized before moving to the next question.
Answer key must be given to the participants after the processing of the case study.
Case studies can be presented in different format. They can be based on the presentation
of a real patient, the files of a patient, a written description of a case, an illustration such
as a photograph or slides of a case, or a video.
This method is not used in this curriculum but trainers can develop case studies based
on local conditions/data as additional exercises if time permits.
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Discussion
Discussion is indicated when the outcome is not predetermined in advance and is “still
negotiable.” Therefore, using discussion to provide “scientific” knowledge/information
or a decision that has already been made and not to be changed can lead to frustration.
Discussion is recommended for plenary or small groups to explore attitudes, values and
opinions. It can confront/compare different options of “doing things” ensuring that the
“why” is covered.
The trainer’s role during the discussion is to facilitate the process, and ensure that the
discussion remains “on track” and that every participant gets a chance to contribute.
When small groups do not have the same assignment/topic to discuss, each group
presents their output(s) and discussion follows immediately after the presentation
before moving to the next group. Time management is essential to ensure that no groups
are “short changed” and has ample time for the presentation and discussion.
If all the groups have the same assignment, all groups present before discussion takes
place. Only clarification questions are allowed during the presentation. Processing the
output(s) must focus on the points of agreement before moving to the differences.
If time does not allow for all groups to present, one group can present and the other
groups complete from their own group’s output before the discussion starts.
Every discussion must be followed by a summary.
Demonstration
Demonstration is a very effective way to facilitate learning of a skill or initiation of the
development of an attitude. The facilitator should use this method to show the skill(s)
and/or the attitude(s) addressing more than one sense at a time. Often a demonstration
can effectively replace a presentation provided the facilitator explains as s/he is doing.
A demonstration should always be followed by a Q&A clarification session before the
learners are required to do a return demonstration.
Steps for a Demonstration
1.
2.
3.
4.
Trainer assembles resources needed for the demonstration
Trainer ensures that participants are ready, can hear and see
Trainer explains what s/he is going to do
Trainer instructs participants on what is expected of them (e.g. to observe closely,
to take notes if appropriate, to use the skills checklist when appropriate etc.)
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5.
6.
 To prepare for the Q&A, and
 Because they are required to do return demonstration(s) for practice
Trainer demonstrates while explaining the skill(s)/attitude necessary for each step
of the procedure being demonstrated
Trainer conducts a Q&A for clarification at the end of the demonstration
Return Demonstration
Return demonstrations provide the learners the opportunity to practice the skills
necessary to perform the procedures they are being trained on. The trainer must ensure
that each learner/participant has the opportunity to practice enough times to reach a
preset minimum acceptable level of performance.
Steps for a Return Demonstration
1. Trainer reminds participants of what is expected of them:
 To practice the procedure/skills
 To observe when others are practicing to be able to ask for clarification
 To observe when others are practicing to be able to provide feedback and peer
evaluation
2. Trainer divides participants into small groups, if more than one workstation.
(Note: each workstation requires at least one facilitator/trainer).
3. Participants take turns practicing the procedure/skills
4. Trainer ensures that all participants can hear and see
5. While each participant is practicing trainer can provide guidance as necessary
provided it does not interfere with the process and confuse the participant
6. After each participant, trainer solicits feedback from other participants
7. After feedback from other participants, trainer reinforces what is correct and
corrects and/or completes feedback
8. Each participant needs to practice more than once or until control of the skill, as
time permits
9. If participant(s) need more than time permits, trainer arranges for additional
practice opportunities
Simulation/Simulated Practice
A simulated practice is an effective method to allow participants to practice
procedures/skills in a specific environment. It recreates as closely as possible the “real
world” without the stress involved in practicing procedures/skills that they do not
control yet in the field. It is recommended to have participants practice on models
before they do perform the procedure/ use the skill in the work place. During a
simulation the participant practices tasks that are part of her/his actual role in the
workplace or that s/he will perform in the job s/he is being trained for.
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Use the same steps as for a demonstration/return demonstration practice.
Role – play
Role-plays are an effective method to practice procedures/skills in the training room.
They are especially effective to practice procedures/skills that deal with human
interactions such as health education and counseling sessions. They are also very
effective when the learning objective deal with attitudes.
In a role play participants “play roles” that are not necessarily their roles in the “real
world.” Often they are asked to play the role of someone they would be dealing with.
In this case it is called “role reversal” or “reverse role play”. This allows the participants
to explore and discover how other perceive/live the interaction.
A role play must always be processed to analyze the lessons learned.
Summary
Every time a training method allows for inputs through exchanges between the
trainer(s) and the participants and between the participants themselves; it must be
followed by a summary session to “tie up the loose ends” and provide the participants
with clear answers. If this does not happen, there is the likelihood that the participants
will forget the “correct” answers.
The trainer should facilitate a summary to ensure that there are “no loose ends.” If time
permits, it is recommended to use the summary for evaluation. In this case, the trainer
can use the Q&A method.
Steps for a Summary for Evaluation
1. Trainer asks a participant to summarize
2. Trainers reinforces if the summary is correct/complete
3. Trainer asks another participant to correct/complete if the summary is
incorrect/incomplete
4. Trainer repeats steps 2 and 3
5. Trainer corrects/completes if after 2 or 3 trials the summary is still
incorrect/incomplete.
Discussion Lecture
Discussion Lecture is introducing scientific material to the listeners. It involves them
in the discussion and exchanging viewpoints, raising questions and answering them.
This leads to enriching the training process and increasing the chances of its success.
The main difference between it and the short lecture is that the trainees are given the
chance for questioning and discussion during the lecturing.
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Discussion lecture uses the principles of the lecture and discussion together in applying
this method.
Privileges of the discussion lecture:
1. Drawing the trainees attention because it is a method of communication between
the two sides in more than one direction
2. Increasing the interaction between the trainees and trainer and among trainees
themselves
3. Allowing the exchange of viewpoints
4. Operating according to the rules and principles of seniors education
5. Allowing the provision of information and decision taking in the same session
Faults of discussion lecture:
1. Discussion may lead to the deviation from the basic subject and this neglecting
the fundamental points of the subjects
2. It cannot be used in gaining the skills
3. It may lead to open the door of the discussion about information and firm
decisions that cannot be changed and this leads to disappointment
Evaluation
Evaluation of learning and training objectives
Evaluation or assessment of learning and of training objectives allows trainers, program
managers and participants to know how successful a training program has been. Ongoing evaluation and assessment allows trainers to identify gaps in learning and to fill
those gaps. Evaluation also assists in revising learning experiences for later trainings.
Many strategies can be used to evaluate learning. Some of the most useful methods
include:

Knowledge assessments: Written or oral questions that require
participants to recall, analyze, synthesize, organize or apply
information to solve a problem. The knowledge component of a skill
objective should be assessed prior to beginning skill practice in a
training room or clinical session.

Questionnaires: Written exercises that assist trainers and participants
to identify and fill gaps in knowledge. Questionnaires can be
administered as self-assessments. In some situations, it may be
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reasonable to have participants use course materials or to work
together on questionnaires.

Skill checklists: Observation of a participant performing a skill and
assessment of the performance using a checklist. Simulated practice
(using real items or models in a situation that is similar to actual
practice) should ideally be assessed prior to beginning clinical
practice with clients. The trainer and other participants should use
the checklists to observe simulated (training room) performance and
actual practice and provide feedback to help improve the
performance. The checklists can also be used by the participant for
self-assessment. During the training participants should be trained
on how to use the checklists. They should also be encouraged to use
them after the training to continue assessing their own performance
and improving it.
Additional techniques for evaluation include: projects, reports, daily reflection, on-site
observation, field performance, and discussion.
Each training module includes assessment of learning methods and tools:

Q&A should be used to frequently identify gaps in knowledge and fill them.

Questionnaires are included with every module and can be used for self-assessment.
To use them as self-assessment, participants complete the questionnaire and then
use any course materials to check their own answers. Trainers should work with
participants filling out the questionnaires to make sure that all gaps in knowledge
are filled before practicing and evaluating skills. When time permits, process
responses in plenary to address any issues and fill the gaps in knowledge. At the
end of this activity the answer key needs to be distributed to the participants.

Skills Checklists are included for each of the skills that are included in this training
curriculum. Participants can use the Skills Checklists as learning guides during
practice sessions in training room or clinical sessions. To evaluate skills, trainers
should generally observe participants three times with coaching as needed to ensure
the skills are learned.
Evaluation of the Participants
Participants’ learning can be evaluated through different methods. This includes Q&A,
synthesis of sessions done by selected participants, self-assessment following the
micro-sessions, peer assessment through feedback provided by other participants
following the micro-sessions and assessment of performance by facilitators.
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Each participant will practice more than once, preferably three times. The curriculum
is for use to plan, organize, conduct and evaluate the training through simulated microsessions. A checklist will be used both by participants for self and peer assessment, and
by the facilitators.
Videotaping the micro-sessions or at least significant segments of the micro-sessions
and reviewing the taped segments after each session will enable the participants to
assess their own progress in terms of acquisition of training/facilitation skills. This
evaluation approach is time consuming, but is very effective in helping participants
assess their own performance. It can also stabilize feedback received from their peers
and the trainers/facilitators.
Post – training evaluation of the learners must be conducted within three to six months
after the end of the training. Further post – training evaluation and follow- up can be
integrated into routine supervision. It is highly recommended to use the skills checklists
used during the training for post – training evaluation and follow-up.
Evaluation of the Training
The “End of Training” evaluation can be done through a questionnaire (Form 1). The
participants are asked to respond and express their opinions about various aspects of
the workshop, such as organization, the process, the facilitation, and a general
assessment.
The “End of Module” evaluation can be done through a questionnaire (Form 2). The
participants are asked to respond and express their opinions about various aspects of
the module. This includes the relevance of the module objective to the course ones, the
relevance of the content to the objectives, the adequacy of the content, the presentation
of the content, the effectiveness of the methodology, the facilitation and the sequencing
of the content.
A confidence/satisfaction index can be calculated to determine how confident the
learners feel that they acquired the knowledge and skills necessary to perform the tasks
they have been trained for, and how committed they feel to using those skills to ensure
the quality of the services they are to provide. The confidence index applies to the
training objectives and acquisition of skills and knowledge and to the degree to which
the participants feel that they able to apply what they have learned during the training.
The satisfaction index applies to the organization and implementation of the training.
The items are labeled in the form of statements followed by a scale 5 (Strongly Agree),
4 (Agree), 2 (Disagree), and 1 (Strongly Disagree). Five (5) represents the highest level
of satisfaction/confidence (agreement with the statement) and one (1) represents the
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lowest. The participants are asked to select the level that expressed their opinion best.
A space for comments is provided after each statement.
The confidence and satisfaction indices are calculated by multiplying the number of
respondents by the correspondent coefficient in the scale, then adding the total. The
total is multiplied by 100. The product is divided by the total number of respondents to
the statement multiplied by 5. 60% represents the minimal acceptable level and 80% a
very satisfactory level of performance.
For example, if the total number of respondents is 19 and 7 of them selected 5 on the
scale, 6 selected 4, 4 selected 2, and 2 selected 1, then the index will be
(5x7)+(4x6)+(2x4)+(1x2) multiplied by 100, divided by (5X19). A 100% index would
if the total number of respondents selected 5. In this case it would be 95. In this example
the index is 72.63%.
The training content and process are evaluated on a continuing basis through daily
evaluations using methods such as “things liked the best” and “things liked the least”
and/or “quick feedback” forms. The facilitators will use the results of this evaluation
during their daily meeting to integrate the feedback and adapt the training to the
participants needs.
“Where Are We?” sessions will be conducted with the participants to assess the
progress in content coverage and process towards reaching the training goals and
learning objectives.
Comments are analyzed and categorized. Only significant comments, those mentioned
more than once and/or by more than one participant, are retained. The facilitators need
to use the results of this evaluation during their daily meeting to integrate the feedback
and adapt the training to the participants needs. Feedback and assessment of training
experiences allows trainers and program managers to adapt training to better meet
participants’ needs. Trainers can also assess their own performance in facilitating the
learning experience of participants using a standardized “facilitation skills” checklist
(Form 4).
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Form 1: END OF COURSE EVALUATION QUESTIONNAIRE
TRAINING CENTER:
DATE:
COURSE TITLE:
INSTRUCTIONS
This evaluation will help adapt the course to your needs and to those of future
participants.
It is anonymous. Please respond freely and sincerely to each item. The items are labeled
in the form of statements followed by a scale where:


5 = strong agree
4 = agree


2 = disagree
1 = strongly disagree
Please circle the number that expresses your opinion; the difference between strongly
agree and agree, and between strongly disagree and disagree are a matter of intensity.
Add your comments in a specific and concise manner, in the space provided after each
statement. If that is not sufficient feel free to use extra paper. If you select 2 or 1, make
sure to suggest how to make the situation better, practical, and offer solutions.
N.B:
Course goals objectives and duration will vary based on the type of training
conducted. Adapt the form to each specific course by including in it the relevant
course items.
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COURSE GOALS
The Course Achieved Its Goals
1. To provide the participants with the opportunities to acquire/update the knowledge
and skills necessary to:
1.1 Play an effective role as a member of the Primary Health Care (PHC)
Center team to improve the quality of care and services
5–4–2–1
Comments:
1.2 Use the team approach to solve problems at the PHC center level
5–4–2–1
Comments:
2. Provide the participants with opportunities to be exposed to and
5–4–2–1
initiate the development of attitudes favorable to the systematic use of
the knowledge and skills acquired in team building and problem solving
to improve the quality of care and services
Comments:
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COURSE OBJECTIVES
1. The course helped me reach the stated objectives:
1.1 Apply the team approach principles to play an effective role as a
member of the Model PHC Center service delivery team
5–4–2–1
Comments:
1.2 Use the team approach to implement the problem solving cycle to
solve service delivery and management problems at the PHC Center
level
5–4–2–1
Comments:
1.3 Explain the importance of being an effective team member of the
Model PHC Center to improve the quality of care and services
5–4–2–1
Comments:
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1.4 Explain the importance of using the team approach to implement the
problem solving cycle to solve service delivery and management
problems at the Model PHC center
5–4–2–1
Comments:
2. The course objectives are relevant to my job description / task I perform in my job:
5–4–2–1
Comments:
3. There is a logical sequence to the units that facilitates learning: 5 – 4 – 2 – 1
Comment:
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ORGANIZATION AND CONDUCT OF THE COURSE
1. Time of notification was adequate to prepare for the course
5–4–2–1
Comments:
2. Information provided about the course before arriving was adequate
5–4–2–1
Comments:
3. Transportation arrangements during the course were adequate (if 5 – 4 – 2 – 1
applicable)
Comments:
4. Training site (Training Center) was adequate
5–4–2–1
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Comments:
5. The educational materials (including reference material) used were 5 – 4 – 2 – 1
adequate both in terms and quantity and quality in relation to the
training objectives and content
Comments:
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6. The methodology and technique used to conduct the training were effective in
assisting you to reach the course objectives
5–4–2–1
7. Clinic/ practice site, as applicable, was adequate
5–4–2–1
Comments:
8. Relationships between participants and course managers and support 5 – 4 – 2 – 1
staff were satisfactory
Comments:
9. Relationships between participants and trainers were satisfactory and 5 – 4 – 2 – 1
beneficial to learning
Comments:
10. Relationships between participants were satisfactory
5–4–2–1
Comments:
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11. The organization of the course was adequate (Time, breaks, supplies,
resource materials)
5–4–2–1
Comments:
Additional comments:
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GENERAL ASSESSMENT
1. I can replicate this training in my future work
5–4–2–1
Comments:
2. I would recommend this training course to others
5–4–2–1
Why or Why Not?
3. The duration of the course (10 days) was adequate to reach all
objectives and cover all necessary topics
5–4–2–1
Comments:
General comments and suggestions to improve the course (Please be specific)
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Form 2: END OF MODULE EVALUATION QUESTIONNAIRE
COURSE: DATE:
MODULE NUMBER & TITLE:
INSTRUCTIONS
This evaluation is intended to solicit your opinions about the modules.
Your feedback will help adapt the course to your needs and to those of future
participants.
It is anonymous. Please respond freely and sincerely to each item. The items are labeled
in the form of statements followed by a scale where:


5 = strongly agree
4 = agree


2 = disagree
1 = strongly disagree
Please circle the number that best expresses your opinion; the differences between
strongly agree and agree, and between strongly disagree and disagree are a matter of
intensity.
Add your comments in a specific and concise manner in the space provided after each
statement. If that space is not sufficient feel to use extra paper. If you select 2 or 1,
make sure to write specific comments on how to improve the module.
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EVALUATION ITEMS
1. The module objectives are relevant to the course objectives
5–4–2–1
Comments:
2. The content / topics covered in the unit are relevant to the 5 – 4 – 2 – 1
objectives
Comments:
3. The content / topics were adequate to help me achieve the 5 – 4 – 2 – 1
objectives
Comments:
4. The content / topics were clear and well-presented
Comments:
5–4–2–1
5. The training methods and activities were effective in facilitating 5 – 4 – 2 – 1
learning
Comments:
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6. The training methods and activities were conducted adequately to 5 – 4 – 2 – 1
facilitate learning
Comments:
7. These are important topics that will enable me to better perform my 5 – 4 – 2 – 1
job
Comments: (specify these points)
8. There is a logical sequence to the sessions and topics that facilitates 5 – 4 – 2 – 1
learning
Comments:
9. There are certain topics that need further clarification
Comments: (specify these points)
5–4–2–1
10. The training materials and resources provided were adequate
Comments:
5–4–2–1
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11. Training materials and resources were provided on time to facilitate 5 – 4 – 2 – 1
learning
Comments:
12. The training materials and resources used were adequate to facilitate 5 – 4 – 2 – 1
my learning
Comments:
13. The training site was adequate
Comments:
5–4–2–1
14. The clinic/ practice site was adequate (if applicable)
Comments:
5–4–2–1
General comments (if any not covered):
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Form 3: QUICK FEEDBACK FORM
TRANING COURSE: DATE:
LOCATION:
MODULE NUMBER AND TITLE:
SESSION NUMBER AND TITLE:
INSTRUCTIONS
This evaluation is anonymous. Please respond freely and sincerely to each item. The
items are labeled in the form of statements followed by a scale where:
5 = strongly agree
4 = agree
2 = disagree
1 = strongly disagree
Please circle the description that expresses your opinion best; the difference between
strongly agree and agree, and between strongly disagree and disagree are a matter of
intensity.
Add your comments in a specific and concise manner, if you have any, in the space
provided after each statement. If that space is not sufficient feel free to use extra paper.
If you selected 2 or 1 please make sure to give comments (e.g. Why? Solutions?)
1. The session objectives are relevant to the tasks in the job description
5–4–2–1
Comments:
2. The methods/learning activities were adapted to the objectives
5–4–2–1
Comments:
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3. The materials provided were adequate to cover all of the content
5–4–2–1
Comments:
4. The time allocated to the session was adequate to cover all the topics
5–4–2–1
Comments:
5. The facilitation (conduct of the session) helped reach the session 5 – 4 – 2 – 1
objectives
Comments:
6. The content of the training was clearly presented
5–4–2–1
Comments:
7. The materials/resources were used in a way that helped me learn
5–4–2–1
Comments:
8. There are points of content that need further clarifications
5–4–2–1
(Specify what specific content areas)
Other Comments:
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Form 4: TRAINING SKILLS CHECKLIST
This checklist is used with the relevant curriculum to give feedback on the trainer’s
performance.
The checklist contains a list of items to be observed:



If they are observed a check mark (√) is entered in the column observed under
adequate or inadequate depending on the performance.
Comments are entered in the appropriate column to clarify/specify what is observed
or not observed.
Is not observed a check mark (√) and comments are entered in the appropriate
columns.
The finding and comments are analyzed and discussed with the trainers supervised. Any
immediate corrective action(s) taken and further action(s) needed must be entered in
the spaces provided.
The trainers supervised must be given an opportunity to comment and the comments
must be entered in the appropriate space. The form must be dated and signed by the
trainer and the supervisor. It is then filed in the trainer’s file for future follow-up and
reference.
Legend: A = Adequate
NA = NOT adequate
Items
Observed
A
NO = NOT observed
NA
NO
Comments
1.Planning of the session
 Relevant sessions plan selected from
curriculum
 Organization conduct and evaluation of
training in conformity with curriculum
(based on observation during the
session)
2.Organizing the session
 Arrive before beginning of session
 Ensure that all training resources are in
place
 Ensure that equipment is in working
condition
 Ensure that training site is set up in
accordance with the requirements of
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the training objective (s) and
methodology
 Prepared/rehearsed for the training
(based on observation of mastery in
conducting activities and using
resources during training)
Items
Observed
A
NA
NO
Comments
3. Conducting the session
3.1 Introduction
 Introduce oneself
- Name
- Job
- Experience relevant to topic
 Introduce/let team members
introduce themselves
 Module:
- Introduce topic
- Present objective
- Clarify topic and objectives
- List sessions
- Establish linkage with
job/task
 Session
- Introduce topic
- Present objectives
- Clarify topics and objectives
- Establish linkage with module
- Establish linkage with
preceding session(s)
- Explain methodology
 Present evaluation methodology
 State estimated duration
3.2 Facilitation skills
 Clarifying
 Make sure participants are ready
before starting on any content
item
 Make sure participants can hear:
- Trainer
- Other participants
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 Make sure participants can see:
- Writing
- Illustrations/ educational aids
- Trainer
- Each other
 Make sure s/he look at
participants
 Make sure s/he can hear
participants
 Use appropriate educational
material
 Summarize after each content
topic item before moving to next
topic
 Use examples relevant to
objectives, content, and
participants learning.
Items
Observed
A
NA
NO
Comments
 Ensuring Active
Participation
 Ask participants questions
 Allow participants to ask
questions
 Allow participants to
question/discuss/make
contributions
 Ensure that all participants
contribute
 Provide participants with
opportunities to practice
 Adapt to participants’ learning
capability (speed, learning
activities, use of educational
material)
 Encourage participants through:
- Listening
- Letting participants complete
their interventions
- Not being judgmental
- Maintaining cordial
relationships with participants
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 Mastering Training
 Conduct the learning activities as
per session plan
 Use the training resources/
materials as per plan
 Cover content adequately
(relevant, clear, concise,
complete, concrete, credible,
consistent and correct)
 Follow curriculum for
learning/training activities
 Use content as per curriculum
1. Evaluating learning/training
process
 Check that participants
understand
 Check that participants learn
skills
 Provide supportive feedback by:
- Reinforcing the positive
learning
- Correcting any errors
- Correcting any incomplete
learning
 Listen to participants comment
about one’s performance (without
making it personal)
 Adapt one’s performance based
on feedback from participants
 Allow participants to answer
questions asked by the group.
Additional comments or observations
Analysis of findings
30 | P a g e
Action (s) taken
Further action (s) needed
Trainer’s comments
Trainer’s name & signature
Supervisor’s name & signature
Date:
31 | P a g e
Part Two
Training Modules
32 | P a g e
Module One: Introduction, Setting up a primary health care
laboratory, and safety in the laboratory
Module Objectives:
At the end of this module the participant will be able to:
1. Identify Setting up a primary health care laboratory
2. Explain safety in the laboratory
Modules Sessions


Session 1: : Introduction and Setting up a primary health care laboratory
Session 2: safety in the laboratory
Evaluation/ Assessment
Questions and answers, participants’ summaries, trainer’s evaluation
Estimated Training Time
3.30 hours
33 | P a g e
Module 1
Session 1: Introduction and Setting up a primary health care laboratory
Objectives
At the end of this session participants will be able to:
1. Explain introduction about laboratory standards
2. Identify Setting up a primary health care laboratory
Trainers Preparation:




Review the reading material and the session plan.
Prepare the presentation as appropriate and as recommended in the method
column of the session plan, or write the information on a flipchart or board
where all participants can see it.
Prepare copies of the reference materials/handouts and exercises.
Arrange the training room.
Methods and activities

Mini-lecture
Evaluation/assessment
Questions and answers, trainer’s observation
Estimated Time
50 minutes
34 | P a g e
Session plan -1-
35 | P a g e
Objectives
1.1.1. Explain
introduction about
laboratory standards
Content


1.1.2.Identify Setting up a
primary health care
laboratory
The "Laboratory
Standard Operating
Procedures," often
called the
"Methods,"
discusses how the
experiment
occurred
Documenting the
procedures of the
laboratory
experiment is
important not only
so that others can
repeat the results
but also so that you
can replicate the
work later, if the
need arises
Methodology
Mini-lecture
25 minutes
Mini-lecture

Figure 1: Plan for a
one room
laboratory
25 minutes
36 | P a g e
Setting up a primary health care laboratory

A one room laboratory: Figure 1 sets out the possible arrangement of a
peripheral medical laboratory attached to a health center. It shows a
laboratory suitable for carrying out some or all of the techniques described in
the manual. The plan is limited to one room, since often this is all the space
that is available for the laboratory. The room should measure at least 5m X
6m.
Figure 1: Plan for a one room laboratory
Alternative plan for a one-room laboratory ( check figure2) 1: outpatient’s table; 2:
hand-operated centrifuge; 3: microscopes; 4: hematology area; 5: colorimeter; 6:
water bath ; 7: electric centrifuge; 8: syphilis serology and biochemistry area; 9:
reagent refrigerator; 10: reagent shelf; 11: glassware shelf; 12: balance; 13: staining
box; 14: area for examination of sputum specimens; 15: Bunsen burner; 16: sinks; 17:
waste sink; 18: bed for patients; 19: record-keeping area; 20: area for examination of
stool specimens; 21: area for examination of urine specimens; 22: area for reception
of specimens; 23: gas bottle.
37 | P a g e
Figure 2: Indicates another possible arrangement of a peripheral
laboratory.
38 | P a g e
Module 1
Session 2: Safety in the laboratory
Objectives
At the end of this session participants will be able to:
1. Identify physical, chemical, and biological hazards
2. Explain safety measures
3. Explain Cleaning, disinfection, sterilization and disposal of biohazard waste
4. Explain Disposal of Biohazard Waste
Trainers Preparation:




Review the reading material and the session plan.
Prepare the presentation as appropriate and as recommended in the method
column of the session plan, or write the information on a flipchart or board
where all participants can see it.
Prepare copies of the reference materials/handouts and exercises.
Arrange the training room.
Methods and activities

Question and answers, discussion-lecture, mini-lecture, discussion
Evaluation/assessment
Questions and answers, trainer’s observation
Estimated Time
150 minutes
39 | P a g e
Session plan 2
Objectives
Content
1.2.1. Identify physical,
chemical, and biological
hazards

1.2.2. Explain safety
measures











Methodology
Sources of
laboratory hazards :
- Physical
- Chemical
- Biological
Questions & answers
Safely Designed
and Organized
Laboratory
Safe use of
Laboratory
Equipment
Safe use of
Electrical
Equipment
Safe use and
Storage of
Chemicals and
Reagents
Preventing
Laboratory
Infection
Practice of Personal
Hygiene
Strict Prohibition of
Mouth Pipetting
Miscellaneous
Elements of a
Laboratory Safety
Program
Management
Responsibility
Safety Officer
Responsibility
Discussion- lectures
30 minutes
60 minutes
40 | P a g e


1.2.3. Explain Cleaning,
disinfection, sterilization
and disposal of biohazard
waste
1.2.4. Explain Disposal
of Biohazard Waste
Supervisor
Responsibility
Employees’
Responsibility



Cleaning glassware
Glass containers
Pipettes





General rules
Laboratory waste
Sharps
Residual body fluids
Chemical wastes
Mini-lecture
30 minutes
Discussion
30 minutes
41 | P a g e
Safety in the Laboratory
Quite common hazards and accidents occur in the laboratory, the need and importance
of laboratory safety should be the real concern of the laboratory. Laboratory accidents
and hazards are controlled by the use of:
 Simple precautions
 Foresight (prudence)
 Safety devices
Above all a ‘real concern’ or a ‘built in concern’ or ‘safety mindedness’ for oneself
and the other fellow working at the next bench is very essential.
Laboratory accidents range from minor injury, illness or loss of body parts to death.
There are many factors that contribute to laboratory accidents. These include:
 Poorly designed laboratory
 Overcrowding of materials
 Poor training
 Lack of concentration
 Noisy and untidy working environment
 Carelessness and neglect
 Overwork and fatigue
 Hot and humid climatic conditions
 Hurrying to finish work on time
 Emergency condition (especially during night hours)
Source of Laboratory Hazards
 Physical
 Chemical
 Biological
1. Physical Hazards
A. Poorly Designed Laboratory Buildings
Due attention should be given in the design of laboratories.
The management, the laboratory personnel and the architecture should be involved in
the standard design of the lab,
B. Burns
Burns may be caused by:
 Flammable chemicals and stains, or by reagents catching alight.
 Fires from spirit lamps, Bunsen burners, lighted tapers (e.g. when heating
Ziehl-Neelsen stain), or from faulty electrical equipment or overloaded
circuits. Spirit burners should not be used in direct sunlight because in bright
light the flame can be difficult to see.
42 | P a g e

Corrosive chemicals being spilt on the skin or ingested when mouth –
pipetting.
C. Electric shock
Electric shock can be caused by:
 Faulty electrical circuits.
 Incorrect installation of equipment.
 Touching exposed live wires.
D. Cuts
Cuts may be caused by:
 Breakages.
 Using glassware that is cracked or has damaged edges.
 Walking on glass chippings.
2. Chemical Hazards
A. Toxic harmful chemicals
Inhaling fumes from toxic chemicals
Ingesting toxic chemicals by mouth pipetting
Skin contact
B. Explosive chemicals
Injury from explosions can be caused by:
Incompatible chemical exploding
Leaking gas
C. Flammable chemicals causing fire
Burns
D. Kinds of chemicals
a. Corrosive (strong acids & alkalis)
 Concentrated sulphuric acid
 Nitric acid
 Sodium hydroxide
 Potassium Hydroxide
b. Toxic irritating chemicals-cause death or serious ill
health if swallowed, inhaled, and by skin contact.
 Potassium cyanide – Chloroform
 Barium chloride - Sodium azide
c. Flammable chemicals
 Ether – Acetone
 Romanowsky stains - Methanol
d. Explosive chemicals
 Picric acid
e. Carcinogens
 Chemicals that cause cancer through ingestion, inhalation, skin contact
 Proven carcinogen chemicals include benzidin, otoulidine, and Nitrophenol.
43 | P a g e

The risk is proportional to the length of exposure, frequency of exposure, and
concentration of the chemical.
3. Biological Hazards
Laboratory acquired infections
Infection can be caused by:
 Pathogens being inhaled in aerosols (airborne droplets) when snap-closing
specimen containers, dispensing or pipetting infectious fluids, or centrifuging
infectious material in open buckets. Aerosols may also be formed and inhaled
following breakages or after spilling infectious fluids. Breakages in
centrifuges can be particularly hazardous if the centrifuge is opened before the
aerosols have settled.
 Pathogen being ingested from contaminated fingers, or in food that has been
contaminated, e.g. by being stored in a laboratory refrigerator. Care should be
taken to avoid the fingers or other parts of the body touching infected material.
Mouth-pipetting specimens and cultures is one of the commonest ways of
ingesting pathogens.
 Pathogens entering the skin through needle punctures, cuts, scratches, insect
bites, sores or other open skin lesions. Laboratory workers must always handle
infected needles with great care.
 Pathogens can also be acquired from unclean or nondisinfected room floors
and walls, water taps and laboratory benches.
 Pathogens are acquired directly through careless contacts with patients or
carrier staff through breathing, hand contact, etc.
4 .Safety Measures
Safely Designed and Organized Laboratory
It is clear that a poorly designed laboratory and overcrowding can increase the risk of
laboratory accidents. It is, therefore, important to know how the laboratory should be
designed with regard to safety considerations. The following are some of the features.
A. Adequate floor, bench and storage space
B. A floor that is well constructed with a surface that is nonslip, impermeable to
liquids, and resistant to those chemicals used in the laboratory.
C. Walls those are smooth, free from cracks, impermeable to liquids and easily
washable.
D. A door at each end of the lab so that lab staff will not be trapped should a
firebreak out.
E. Adequate ventilation with windows that can be opened.
F. Sectioning of the laboratory into separate rooms with places for patients,
visitors, and reception of specimens.
G. Bench surface that are without cracks, impervious, washable, and resistant to
disinfectants and chemicals.
44 | P a g e
H. Suitable storage facilities that include a well-ventilated, fire proof, locked
store, for the storage of flammable chemicals.
I. A good supply of gas, water, electric power & wall electric points
J. Provision of protective safety cabinets, fire extinguishers at accessible points,
and adequate waste disposal area, etc.
Safe use of Laboratory Equipment
A. Positioning
 Suitable and ideal place for operation
 Avoid overcrowding of a bench with equipment
 Position equipment correctly that requires special facilities like ventilation,
shield from sunlight, and great care.
B. Installation

Should be carried out by the supplier or by the health unit electrician or trained
lab equipment technician.
 Important points to consider for safe installation
 Ensure that the voltage of the new equipment is the same as that of the
electricity supply.
 Check that the power required by the instrument does not exceed the power
supply circuit of the lab.
 Make sure that the equipment is wired correctly, and the wiring system have
grounded conductor.
Safe use of Electrical Equipment
The supplier should demonstrate the use of an apparatus. If this is not possible, the
operation and service manual should be carefully studied before the equipment is
operated.
Points to consider with regard to the safe use of electrical equipment:
1. Hands should be dry completely, and also the floor on which the operator is
standing.
2. The electric supply must be disconnected when performing any maintenance
and at the end of the day’s work.
3. If a fuse should blow, do not automatically put in a new one until the circuit is
checked.
Safe use and Storage of Chemicals and Reagents
Even in the smallest lab, dangerous chemicals are used directly or incorporated into
stains and reagents. These include highly flammable chemicals such as ether or
methanol, highly corrosive chemicals such as phenol or sulphuric acid, or toxic and
harmful chemicals such as formaldehyde solution.
The correct handling and storage of hazardous chemicals is essential to prevent injury
and damage. It is particularly important to keep chemicals out of direct sunlight and
avoid overheating in chemical stores and the laboratory.
Overheating can decompose many chemicals, cause explosions, or the formation of
toxic fumes.
45 | P a g e
Labeling of dangerous chemicals and reagents
To reduce accidents caused by chemicals, many countries have introduced legislation,
requiring manufacturers to label dangerous chemicals with hazard symbols and to
provide simple safety instructions.
The six accepted danger symbols currently in use are toxic, corrosive, explosive,
oxidizing, highly flammable, and harmful or irritant.
Figure 3: Common symbols of hazards
Preventing Laboratory Infection
All specimens received in the lab should be regarded as potentially pathogenic. For
example, a blood specimen sent for measuring hemoglobin may contain highly
infectious organisms.
Laboratory acquired infections can be prevented by:
 Practicing personal hygiene
 Wearing of laboratory coat and protective gloves
 Safe handling of specimens and infectious materials.
 Avoiding mouth-pipetting
 Disposing safely of specimens and contaminated material.
 Being immunized against highly infectious pathogens
Practice of Personal Hygiene
This includes:
 Washing of hands and arms with soap and water before and after work
 Wearing protective clothing and gloves whenever possible.
 Covering any cuts, insect bites open sores, or wounds with a water proof
adhesive dressing.
 Wearing closed shoes and not walking barefoot.
 Not eating, drinking, chewing gum, smoking or applying cosmetics in any part
of the lab.
Safe Handling and Disposal of Specimens
Special precautions should be taken when collecting specimens, especially blood
specimens, and when testing specimens and handling infected material.
Safety measures involved are:
 Careful handling and disposal of materials used for collecting specimens.
 Making contaminated materials non-infectious by using appropriate
decontamination methods.
46 | P a g e

Wearing gloves and a plastic apron when collecting blood suspected of having
a highly infectious disease like AIDS.
 Avoid contamination of fingers, other body parts, and working surface.
 Specimens suspected of having hepatitis, and AIDS must be labeled ‘HIGH
RISK’.
Strict Prohibition of Mouth Pipetting
Pathogens may be ingested during mouth pipetting, either by direct aspiration or from
the mouth ends of pipettes which have been contaminated from fingers or benches.
Accidents caused by mouth pipetting include infection, poisoning, chemical burns,
and other injuries from chemicals. There are many inexpensive and simple ways to
measure and dispense safely without mouth pipetting.
Miscellaneous
 Immunization- protective inoculations against certain pathogenic organisms
are necessary.
 The use of signs- display suitable safety signs both prohibitive (don’t) and
commands (do).
 First aid- basic practical training in first aid helps to reduce suffering and
consequences of serious accidents.
Elements of a Laboratory Safety Program
A successful laboratory safety program requires the participation of persons at every
level of the laboratory staff.
 Safety does not occur only with the appointment of safety officer or safety
committee.
 Safety does not occur by having employees solely perform their assigned
duties in manner which they feel is most efficient.
 Laboratory safety requires the full participation of every member of the staff.
Management Responsibility
 Establish a policy relative to the design and implementation of his safety
program.
 Delegate authority for implementing the program
 Provide a safe and healthful work place
 Provide fund for the implementation of the program
 Assess the program- establish a mechanism to ensure safety
 Establish safety committee- regular report, recommendations regarding need
for modification of the program.
Safety Officer Responsibility
 Technical advisor to the program
 Assist in the development of safe work method
 Advise management on safety issues
 Assist safety committee
 Provide a variety of communication, e.g. hazard notice, safety data
47 | P a g e

Review a variety of plans which include facility designs, special equipment
purchase in relation to safety.
Supervisor Responsibility
 Train the staff in lab practices required for safe conduct of work.
 Evaluate regularity of the laboratory facilities, equipment, personnel and work
place.
 Correct unsafe condition as fire hazards, physical hazards, and defective
equipment.
Employees’ Responsibility
 Use of safe equipment
 Report of malfunctioning of equipment
 Report injuries or exposure
 Report hazard or unsafe condition to supervisors

5. Cleaning, disinfection, sterilization and disposal of biohazard waste.
Cleaning glassware
Instructions for cleaning:
 glass containers (Erlenmeyer flasks, beakers, test-tubes)
 pipettes
Glass containers
New glassware
Glassware that has never been used may be slightly alkaline.
In order to neutralize it:

Prepare a bowl containing 3 liters of water and 60 ml of concentrated
hydrochloricacid (i.e. a 2% solution of acid).
 Leave the new glassware completely immersed in this solution for 24 hours.
 Rinse twice with ordinary water and once with demineralized water.
 Dry.
Dirty glassware
 Preliminary rinsing: Rinse twice in cold or lukewarm water (never rinse
bloodstained tubes in hot water), If the glassware has been used for fluids
containing protein, it should be rinsed immediately and then washed (never
allow it to dry before rinsing).
 Soaking in detergent solution: Prepare a bowl of water mixed with washing
powder or liquid detergent. Put the rinsed glassware in the bowl and brush the
inside of the containers with a test-tube brush. Leave to soak for 2–3 hours.
Rinsing
Remove the articles one by one. Rinse each one thoroughly under the tap, then soak
them all in a bowl of ordinary water for 30 minutes.
48 | P a g e
Rinse each article in a stream of clean water. (Do not forget that traces of detergent
left on glassware can lead to false laboratory results.)
Draining
Place containers (beakers, flasks, measuring cylinders) on the pegs of a draining rack.
Place test-tubes upside-down in a wire basket.
Drying
Place the glassware in wire baskets and dry in a hot-air oven at 60 °C. Alternatively,
place the baskets in a sunny spot in the laboratory and cover them with a fine cloth.
Plugging
The clean dry glassware should be put away in a cupboard to protect it from dust. It is
recommended that glass containers be plugged with non-absorbent cotton wool or
their mouths covered with small caps made from newspaper or, preferably, thin sheets
of paraffin wax or clinging plastic, if available.
Pipettes
Immediate rinsing
Once a pipette has been used, rinse it immediately in a stream of cold water to remove
blood, urine, serum, reagents, etc.
Soaking in water
After rinsing, place the pipettes in a large, plastic measuring cylinder (or bowl) full of
water. If the pipettes have been used to measure infected material, leave them in a
cylinder full of disinfectant solution (e.g. a quaternary ammonium compound or
1% bleach solution; see pages 84 and 85) for 4 hours.
Soaking in detergent and rinsing
Follow the instructions given above for soaking and rinsing of laboratory glassware.
Blocked pipettes
Put blocked pipettes in a cylinder filled with dichromate cleaning solution. Slide them
carefully into the solution and leave for 24 hours.
The next day, pour the dichromate solution into another cylinder (it can be used four
times).
Hold the cylinder containing the pipettes under the tap and rinse thoroughly.
Remove the pipettes one at a time. Check that the obstruction has been washed away.
Rinse again.
Leave to soak in ordinary water for 30 minutes, then change the water and soak for a
further 30 minutes.
Warning: Dichromate cleaning solution is highly corrosive and should be used with
extreme care. If it is accidentally splashed on the skin or clothing or into the eye(s),
wash at once with large quantities of water.
49 | P a g e
Drying
Dry heat-resistant glass pipettes in a hot-air oven at 60 °C and ordinary pipettes in an
incubator at 37 °C. Alternatively, leave pipettes to air-dry.
Using the vacuum pump
This is a small instrument made of metal, plastic or glass that is attached to the water
tap.
Turn the water on hard to drive a strong jet through the pump. This causes air to be
sucked into the side arm of the pump and the rubber tubing attached to it.
Fit this rubber tubing over the tip of the pipette.
3. Dip the other end of the pipette into the rinsing liquid (water or detergent solution),
which is sucked through the pipette and discharged by the pump into the sink (Fig ).
6. Disposal of Biohazard Waste
A. General rules
Prior to any treatment, all biohazard wastes, including those to be incinerated, should
be enclosed in a puncture-resistant, biohazard bag that is color-coded or labeled with
the biological hazard symbol.
Untreated biohazard waste is not to be disposed of in the municipal waste (system)
stream. All biohazard waste must be treated by chemical disinfection or autoclaving
before in any way being disposed of in the municipal waste stream.
50 | P a g e
After disinfection, but before disposal in the municipal waste stream, all treated
biohazard wastes (apart from liquids) should be enclosed in an unmarked outer bag
that is not red or labelled with the biohazard symbol. Any biohazard waste that has
been treated as described below and packaged such that it is clearly evident that the
waste has been effectively treated, is not subject to be treated as biohazard waste and
may be collected, transported for incineration or disposed of as municipal waste.
The person handling the emptying of waste bins, waste bottles or sharps containers
must be careful not to touch anything without protective clothing and must use gloves
to avoid contracting infections from the waste.
The waste bins, sharps containers, etc., must be clearly indicated/marked:
Biohazard wastes.
B. Laboratory waste
Discarded swabs, cotton, sample containers and other biohazard but non-sharp objects
used for sample collection should be placed in a covered trash container lined with an
autoclavable biohazard yellow bag. Visibly soiled or splashed tourniquets and
Vacutainer holders should be discarded and new equipment used.
Disposable plastics such as tips, culture plates, micro plates, petri dishes, test tubes,
etc., used for testing of samples should be disposed of in a covered waste bin lined
with an autoclavable biohazard yellow bag, autoclaved and transported for
incineration.
Pipettes should be soaked in special pipette baskets in 10 % household bleach
overnight, before being thrown in a waste bin lined with an autoclavable biohazard
yellow bag, autoclaved and incinerated.
Reusable glassware such as cylinders, flasks and beakers should be disinfected with
10 % household bleach over night before washed and autoclaved.
C. Sharps
All sharp implements used in the laboratory need to be handled carefully. They can
easily break the skin and increase the risk of infection with infectious agents.
Broken glassware must also be handled as sharps. The laboratory should have a
special box for broken glassware
If needles are used, they should never be recapped before disposal. Recapping needles
may lead to somebody accidentally pricking him or herself.
Dispose of all sharps directly into a with a secure perforated sharps disposal container
lid. The lid only allows the implements into the container but not out. The sharps
container should be puncture-resistant, leak proof on the sides and bottom, and colorcoded or labeled with a biohazard symbol.
Note: When selecting sharps containers, look for special safety features such as lids
that lock tight for safe disposal and a container that can be sterilized by steam, gas, or
chemicals. If sharps containers are not specifically constructed to be autoclaved, the
51 | P a g e
resulting mass of melted plastic is extremely hazardous due to the needles that often
protrude.
When full, seal top, autoclave, then arrange for incineration.
Decontamination is achieved by autoclaving the sharp implements. Incineration
destroys the sharps completely hence eradicating the chances of accidental injury to
people in the community.
D. Residual body fluids
Ensure that all tubes/containers containing residual body fluids are properly sealed to
avoid spillage. These are then stored temporarily in waste bins lined with disposal
bags.
Transfer the disposal bags, together with contents, into autoclaving bags stick a piece
of autoclavable tape on the bag and then autoclave as required.
After the autoclaving cycle is completed, transport and incinerate the whole package
to completely destroy the tubes.
E. Chemical wastes
The following procedure is adopted for most chemicals used in low concentrations.
Collect all liquid chemical waste in properly labeled bottles with a little concentrated
disinfectant (i.e., JIK).
Keep monitoring the rising level of waste in the trap bottles. Never fill the trap bottles
to the very top to avoid spillage.
Empty the contents of the trap bottle down the drain, preferably a special sink in the
laboratory, and wash down with more disinfectant, liquid soap and a large volume of
water.
Minimum disinfection time of any liquid Biohazard Waste is 30 minutes.
Note:Some disinfectants react with organic compounds, acids, or bases. Consider this
when selecting your disinfectant before commencing your work.
SAFETY PRECAUTIONS
Anybody handling Biohazard Wastes should always put on laboratory protective
clothing and gloves and keep to the laboratory safety practices to avoid viral and other
infectious disease transmissions.
Any spills or splashes of infectious material should be immediately cleaned up with
absorbent material using an approved disinfectant such as dilute 10% bleach or
chlorhexidine solution.
Decontaminating bench tops and other surfaces
Bench tops should be wiped with a disinfectant (a phenolic disinfectant, 1% sodium
hypochlorite [bleach], or 70% isopropyl alcohol) routinely after working with
infectious agents or clinical specimens or after spills, splashes, or other contamination
by infectious materials. Solutions of disinfectants should be maintained at the work
station.
52 | P a g e
Decontamination of spills
The following procedure is recommended for decontaminating spills.
• Isolate the area to prevent anyone from entering.
• Wear gloves and protective clothing (e.g., a gown or lab coat, shoes, and a mask [if
the spill may contain a respiratory agent or if the agent is unknown]).
• Absorb or cover the spill with disposable towels.
• Saturate the towels with an appropriately diluted intermediate or high-level
disinfectant (e.g., a phenolic formulation or household bleach).
• Place disinfectant-soaked towels over the area and leave them in place for at least 15
minutes before removing and discarding them.
• Wipe area using clean disinfectant-soaked towels and allow area to air dry.
• Place all disposable materials used to decontaminate the spill into a biohazard
container.
• Handle the material in the same manner as other infectious waste.
53 | P a g e
Module two: Hematology
Module Objectives:
At the end of this module the participant will be able to:
3. Apply collection and handling of blood
4. Apply Automated Blood Count Techniques
5. Apply manual techniques
Modules Sessions


Session 1: Collection and handling of blood and automated blood count
techniques
Session 2: Manual techniques
Evaluation/ Assessment
Questions and answers, participants’ summaries, trainer’s evaluation
Estimated Training Time
9 hours
54 | P a g e
Module 2
Session 1: Collection and handling of blood and automated blood count
techniques
Objectives
At the end of this session participants will be able to:
1. Apply collection and handling of blood
2. Explain hemoglobin concentration, red blood cell count, and counting systems
3. Explain reliability of electronic counters
4. Explain Packed Cell Volume and Mean Cell Volume
5. Explain variation in red cell volumes: red cell distribution width and total white
blood cell count
6. Explain automated differential count
7. Explain new white cell parameters and platelet count
8. Explain mean platelet volume, reticulocyte count, and immature reticulocyte
fraction
9. Explain calibration of automated blood cell counters
10. Explain flagging of automated blood counts
Trainers Preparation:




Review the reading material and the session plan.
Prepare the presentation as appropriate and as recommended in the method
column of the session plan, or write the information on a flipchart or board
where all participants can see it.
Prepare copies of the reference materials/handouts and exercises.
Arrange the training room.
Methods and activities

Mini-lecture, Question & answers, discussion-lecture, discussion, brain
storming
Evaluation/assessment
Questions and answers, trainer’s observation
Estimated Time
290 minutes
55 | P a g e
Session plan -1Objectives
2.1.1. Apply collection
and handling of blood
Content






2.1.2. Explain
hemoglobin
concentration, red blood
cell count, and counting
systems
2.1.3. Explain reliability
of electronic counters




Methodology
General
precautions:
Technical errors:
Venous blood
collection:
Capillary
(peripheral) blood
Differences
between capillary
and venous blood
Serum
Mini-lecture
Hemoglobin
concentration
Red blood cell
count
Counting systems
- Impedance
counting
- Light scattering
Question & answers
30 minutes
30 minutes
Electronic counts are Mini-lecture
precise, but care
needs to be taken so
30 minutes
that they are also
accurate. The
recorded count on the
same sample may
vary from instrument
to instrument and
even between
different models of
the same instrument.
56 | P a g e

2.1.4. Explain Packed
Cell Volume and Mean
Cell Volume





Setting
discrimination
thresholds
Modern automated
blood cell counters
estimate PCV by
technology that has
little connection
with packing red
cells by
centrifugation
With automated
instruments, the
derivation of the
RBC, PCV, and
MCV are closely
interrelated
Automated
instruments require
calibration before
the PCV or MCV
can be determined
The MCV, and
therefore the PCV,
as determined by
an automated
counter, will vary
with certain cell
characteristics
other than volume
The automated
MCV and PCV are
prone to certain
errors that do not
occur or are less of
a problem with
manual methods
Discussion-lecture
30 minutes
Discussion
2.1.5. Explain variations
in red cell volumes: red
cell distribution width

Automated
instruments
produce volume
30 minutes
57 | P a g e
and total white blood cell
count

distribution
histograms that
allow the presence
of more than one
population of cells
to be appreciated
The total WBC is
determined in
whole blood in
which red cells
have been lysed
Mini-lecture
2.1.6. Explain automated
differential count
30 minutes

2.1.7. Explain new
white cell parameters
and platelet count
Most automated
differential
counters that are
now available use
flow cytometry
incorporated into a
full blood counter
rather than being
stand-alone
differential
counters
Question & answers
30 minutes


Automated white
cell counters can
analyze cell
characteristics by
novel technologies
and identify cell
types by features
that differ greatly
from those used
when a blood film
is examined
visually
. Three techniques
for setting
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thresholds have
been used:
(a) Platelets can be
counted between
two fixed
thresholds (e.g.,
between 2 and 20
fl;
(b) Pulses between
fixed thresholds
can be counted
with subsequent
fitting of a curve
and extrapolation
so that platelets
falling outside the
fixed thresholds are
included in the
computed count.
(c) Thresholds can
vary automatically,
depending on the
characteristics of
individual blood
samples, to make
allowance for
microcytic or
Mini-lecture
fragmented red
30 minute
cells or for giant
platelets
2.1.8. Explain mean
platelet volume,
reticulocyte count, and
immature reticulocyte
fraction


The calculated
MPV is very
dependent on the
technique of
measurement and
on length and
conditions of
storage prior to
testing the blood.
Automated
reticulocyte counts
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
2.1.9. Explain calibration
of automated blood cell
counters

2.1.10. Explain flagging
of automated blood
counts
have been
developed by using
the fact that various
dyes and
fluorochromes
combine with the
RNA of
reticulocytes
Fully automated
instruments provide
a measure of the
various degrees of
reticulocyte
maturation because
the most immature
reticulocytes,
produced when
erythropoietin
levels are high,
have more RNA
and fluoresce more
strongly than the
Mini-lecture
mature
30 minutes
reticulocytes
normally present in
the peripheral
blood
Methods are
recommended for
calibrating an
automated blood
cell counter:
- By using fresh
normal blood
specimens
- By use of a stable
calibrant
- By use of a
commercial
calibrant
Brain storming
20 minutes
60 | P a g e

Flagging” refers to
a signal that the
specimen being
analysed may have
a significant
abnormality
because one or
more of the blood
count variables are
outside specified
limits (usually
2SD) or there is a
qualitative
abnormality that
requires a quality
control check
and/or additional
investigation
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Hematology
1. Collection & Handling of blood
General precautions:




Technical errors:


Always wear disposable plastic gloves, especially if there is any hand cut.
Always be careful when handling syringes, needles & lancets.
Always use disposable syringes, needles & lancets.
Always take care of specimen identification & request forms.
Faults in sample Collection (See below).
Faults in sample Transport & Storage (identification error, how long delay
& at what temp.).
Technical errors from sample collection:

Insufficient or excess anticoagulant.

Inappropriate anticoagulant.

Inadequate mixing with anticoagulant.

Patient &/or specimen identification error.

Delay in transit to lab.
Venous blood collection:
The phlebotomist should enter a training program. Venous blood,
mixed with an anticoagulant, is preferred for most hematological examinations.
Best site is the antecubital vein, & in obese subject may use the dorsum of the hand.
Needle guage: for adults 21 (0.8mm), for children 23 (0.6mm).
Make sure of proximal & distal fixation of the vein.
Skin preferably cleaned with 70% alcohol.
Steps to facilitate obtaining a venous blood sample are:

Warming the patient’s arm.

Using a tourniquet (should be loosened once entered the vein to avoid
hemoconcentration and hemolysis).

Tapping the skin over the site of the vein.

Asking the patient to open & clench his fist several times.
Advantages of using the tourniquet:
- Blood filling of the vein.
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-
Proximal fixation of the vein.
Disadvantages of using the tourniquet:
- Hemoconcentration.
- Red cell hemolysis.
Always notice the blood level marker in tubes containing an anticoagulant.
After puncture: elevate the arm, apply pressure & adhesive dressing.
There is no difference in using glass or plastic tubes except in coagulation study (where plastic or
siliconized glass tubes should be used).
Avoid hemolysis of blood by:

Minimal use of the tourniquet.

Withdraw blood slowly.

Do not use very fine needled.

Deliver the blood gently into the tube.

Avoid frothing during mixing the tube.
Capillary (peripheral) blood:
It is often necessary to obtain blood by skin puncture in babies and infants and in adults with poor
veins.
Capillary blood may be obtained from a freely flowing stab wound made with a sterile lancet on:
- The plantar surface of a warmed and cleansed heel (babies less than 3 months of age). The correct
site for puncture of the heel is shown in Fig. 1.4.
- The plantar aspect of the big toe (infants).
- A finger, thumb or ear lobe (older children and adults).
The lateral or posterior aspect of the heel should not be used in a baby, as the underlying bone is
much closer to the skin surface than it is on the plantar aspect. Osteomyelitis of the calcaneal bone
has resulted from puncture of the bone.
In older patients a finger (excluding the fifth finger) or the thumb is preferred. The palmar surface
of the distal phalanx is the preferred site on a digit, since the underlying bone is closer to the skin
surface on other aspects. The middle or ring finger of the non-dominant hand is preferred; these
digits are less painful than the index finger. Previous puncture sites should be avoided, to reduce
the risk of infection.
In adults, skin punctures should ideally be more than 1.5 mm deep in order that the lancet passes
through the dermal– subcutaneous junction where the concentration of blood vessels is greatest,
permitting a free flow of blood. Lancets used for heel puncture in full-term babies must not exceed
2.4 mm in length, since this is the depth below the skin of the calcaneal bone. Much shorter lancets
are available and should be selected for use in premature babies. Safety lancets, with a blade that
retracts permanently after first use, have been developed in order to reduce the risk of accidental
injury to phlebotomy staff.
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Capillary samples should be obtained from warm tissues so that a free flow of blood is more
readily obtained. If the area is cool then it should be warmed with a wet cloth. The skin should
then be cleansed with 70% alcohol and dried with sterile gauze (since traces of alcohol may lead
to haemolysis of the specimen). The first drop of blood may be diluted with tissue fluid and should
be wiped away with sterile gauze square. Flow of blood may be promoted by gentle pressure, but
a massaging or pumping action should not be employed, since this may lead to tissue fluid being
mixed with blood.
Differences between capillary and venous blood:
Venous blood and “capillary” blood are not quite the same. Blood from a skin puncture is a
mixture of blood from arterioles, veins, and capillaries, and it contains some interstitial and
intracellular fluid.
However, there are negligible differences when a free flow of blood has been obtained. The
differences may be exaggerated by cold with resulting slow capillary blood flow.
The packed cell volume (PCV), RBCs count, and Hemoglobin concentration (Hb) of capillary
blood are slightly greater than in venous blood. Also the total leukocyte and neutrophil counts are
higher.
Conversely, the platelet count appears to be higher in venous than in capillary blood. This may be
the result of adhesion of platelets to the site of the skin puncture.
Serum:
The difference between plasma and serum is that the latter lacks fibrinogen and some of the
coagulation factors.
Blood collected to obtain serum should be delivered into sterile tubes with caps or evacuated
collection tubes and allowed to clot undisturbed for about 1 hour at room temperature (18-25°C).
Then the clot should be loosened gently from the container wall by means of a wooden stick or a
thin plastic or glass rod. Rough handling will cause hemolysis. The tubes are centrifuged for 10
min at about 1200 rpm. The supernatant serum then is pipetted into another tube and centrifuged
again for 10 min at about 1200 rpm. The supernatant serum is transferred to tubes for tests or for
storage.
For most tests, serum should be kept at 4°C until used, but if testing is delayed, serum can be
stored at -20°C for up to 3 months and at -40°C or less for long-term storage. Frozen specimens
should be thawed on the bench or in a water bath at room temperature, and then inverted several
times to ensure homogeneity before use for a test. Do not refreeze thawed specimens.
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Figure 4:Anterior surface of the left arm showing veins most suitable for venepuncture.
Figure 5: Venepuncture technique using needle and syringe.
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Figure 6: Venepuncture technique using an evacuated container; the distal end of the needle
has been screwed into the holder and the proximal needle has then been unsheathed and inserted
into a suitable vein.
Figure 7:The areas of the foot of a baby or infant which are suitable for
obtaining capillary blood.
2. Part One: Automated Blood Count Techniques
A variety of automated instruments for performing blood counts are in widespread
use. semi-automated instruments require some steps (e.g., dilution of a blood sample)
to be carried out by the operator. Fully automated instruments require only that an
appropriate blood sample is presented to the instrument. Semiautomatic instruments
often measure a small number of components (e.g., WBC and Hb). Fully automated
multichannel instruments usually measure from 8 to 20 components, including some
variables that have no equivalent in manual techniques. Automated instruments
usually have a high level of precision, which, for cell counting and cell-sizing
techniques, is greatly superior to that achievable with manual techniques. If
instruments are carefully calibrated and their correct operation is ensured by quality
66 | P a g e
control procedures, they produce test results that are generally accurate. When blood
has abnormal characteristics, the results for one or more parameters may be aberrant;
instruments are designed so that such inconsistent results are “flagged” for
subsequent review. The abnormal characteristics that lead to inaccurate counts vary
between instruments, so it is important for instrument operators to be familiar with the
types of factitious results to which their instruments are prone.
Blood cell counters may have automated procedures for sample recognition (e.g., by
bar-coding), for ensuring that adequate sample mixing occurs, for taking up the test
sample automatically, and for detection of clots or inadequately sized samples.
Ideally, blood sampling is carried out by piercing the cap of a closed tube so that
samples that carry an infection hazard can be handled with maximum safety.
Laboratories performing large numbers of blood counts each day require fully
automated blood counters capable of the rapid production of accurate and precise
blood counts, including platelet counts and differential counts, either three-part or
five- to seven-part. The sample throughput required varies with the workload and the
timing of arrival of blood specimens in the laboratory, but for most large laboratories
a throughput of 100 or more samples per hour is required. Sample size and the
availability of a “predilute” mode are particularly relevant if the laboratory receives
many paediatric specimens.
Choice of an instrument for an individual laboratory, as well as for point-of-care sites
outside the laboratory, should take account of capital expenditure and running costs,
including maintenance and reagents; size of instrument; requirements of services such
as water, compressed air, drainage, and an electricity supply with stable voltage;
environmental disturbance by generation of heat, vibration, and noise; any influence
on performance by the ambient temperature and humidity; storage requirements for
the often bulky reagents; ease of operation; and the likely level of support that can be
expected from the manufacturer.
Most automated instruments, however, count for a specified period of time rather than
on an exact volume of blood; they therefore require calibration by means of the direct
counts derived from instruments counting cells in a defined volume of diluted blood.
For some variables, instruments are calibrated by the manufacturer, but others require
calibration in the laboratory. Performance characteristics of an instrument vary over
time, so periodic recalibration is needed; both when quality control procedures
indicate the necessity and when certain components are replaced.
2.1 Hemoglobin Concentration
Most automated counters measure haemoglobin by a modification of the manual
HiCN method with cyanide reagent or with a nonhazardous chemical such as sodium
lauryl sulphate, which avoids possible environmental hazards from disposal of large
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volumes of cyanide-containing waste. Modifications include alterations in the
concentration of reagents and in the temperature and pH of the reaction. A nonionic
detergent is included to ensure rapid cell lysis and to reduce turbidity caused by cell
membranes and plasma lipids. Measurements of absorbance are made at a set time
interval after mixing of blood and the active reagents but before the reaction are
completed.
2.2 Red Blood CELL Count
Red cells and other blood cells can be counted in systems based on either aperture
impedance or light-scattering technology. Because large numbers of cells can be
counted rapidly, there is a high level of precision. Consequently, electronic counts
have rendered the RBC and the red cell indices derived from it (the MCV and the
MCH) of much greater clinical relevance than was possible when only a slow and
imprecise manual RBC was available.
2.3 Counting Systems
Impedance Counting
Impedance counting, first described by Wallace Coulter in 1956 depends on the fact
that red cells are poor conductors of electricity, whereas certain diluents are good
conductors; this difference forms the basis of the counting systems used in Beckman–
Coulter, Sysmex, Abbott, Roche, and a number of other instruments.
For a cell count, blood is highly diluted in a buffered electrolyte solution. The flow
rate of this diluted sample is controlled by a mercury siphon (as in the original Coulter
system) or by displacement of a tightly fitting piston. This result is a measured
volume of the sample passing through an aperture tube of specific dimensions (e.g.,
100 mm in diameter and 70 mm in length). By means of a constant source of
electricity, a direct current is maintained between two electrodes, one in the sample
beaker or the chamber surrounding the aperture tube and another inside the aperture
tube. As a blood cell is carried through the aperture, it displaces some of the
conducting fluid and increases the electrical resistance. This produces a corresponding
change in potential between the electrodes, which lasts as long as the red cell takes to
pass though the aperture; the height of the pulses produced indicates the volume of the
cells passing through. The pulses can be displayed on an oscillograph screen. The
pulses are led to a threshold circuit provided with an amplitude discriminator for
selecting the minimal pulse height, which will be counted. The height of the pulses is
used to determine the volume of the red cells.
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Figure 8: Effect of threshold discrimination (horizontal axis) in separating cell
signals from background noise.
Light Scattering
Red cells and other blood cells may be counted by means of electro-optical detectors.
A diluted cell suspension flows through an aperture so that the cells pass, in single
file, in front of a light source; light is scattered by the cells passing through the light
beam. The scattered light is detected by a photomultiplier or photodiode, which
converts it into electrical impulses that are accumulated and counted. The amount of
light scattered is proportional to the surface area and therefore the volume of the cell
so that the height of the electrical pulses can be used to estimate the cell volume. The
high-intensity coherent laser beams used in current instruments have superior optical
qualities to the noncoherent tungsten light of earlier instruments. Sheathed flow
allows cells to flow in an axial stream with a diameter not much greater than that of a
red cell; light can be precisely focused on this stream of cells. Electro-optical
detectors are used for red cell sizing and counting in Bayer-Technicon systems and
for white cell differential counting in a number of other instruments.
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2.4 Reliability of Electronic Counters
Electronic counts are precise, but care needs to be taken so that they are also accurate.
The recorded count on the same sample may vary from instrument to instrument and
even between different models of the same instrument. Inaccuracy may be introduced
by :
1-coincidence (i.e., by two cells passing through an orifice simultaneously and
being counted as one cell.
2-pulse being generated during the electronic dead time of the circuit.
3-Recirculation of cells that have already been counted.
4-red cell agglutination (which causes a clump of cells to be counted as one
Cell).
5-by the counting of bubbles, lipid droplets, microorganisms, or extraneous
Particles as cells.
6-Faulty maintenance may lead to variation in the volume aspirated or the flow rate.
7-Single-channel instruments may have their thresholds set incorrectly, and
multichannel instruments may be incorrectly calibrated.
Coincidence correction; in some instruments, this is done automatically by electronic
editing. Errors of coincidence can be detected by carrying out a series of
measurements at various dilutions of the same specimen, plotting the data on graph
paper, and then extrapolating the graph to the baseline for the true value.
Alternatively, the need for coincidence correction can be avoided by having the
dimensions and flow characteristics of the aperture through which the cells pass such
that cells can only pass in single file; this may be achieved by sheath flow or
hydrodynamic focusing in which diluted blood is injected into a sheath of fluid as it
flows into the sensing zone. This induces the cells to pass through the centre of the
sensing zone in single file and free of distortion. Coincidence can be more effectively
reduced with sheathed flow and precisely focused light in an electro-optical detector
than in an impedance counter so that less dilution of the blood sample is needed.
Electrical impulses generated by recirculation of cells can be eliminated by electronic
editing.
recirculation of cells in the region of the aperture can be prevented by “sweep flow”
in which a directed stream of diluent sweeps cells and debris away from the aperture,
thus preventing cells from being recounted and debris from being counted as cells.
Inaccurate counts consequent on red cell agglutination are usually the result of cold
agglutinins. They are recognized as erroneous because of an associated marked
factitious elevation of the MCV. A correct count can be achieved by prewarming the
blood sample and, if necessary, also prewarming the diluent.
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A correct RBC and, particularly, a correct measurement of the MCV are dependent on
the use of an appropriate diluent. For impedance counters, pH, temperature, and rate
of ionization have to be standardized and remain constant because changes alter the
electrical field and may lead to artefactual alterations in the size, shape, and stability
of the blood cells in the diluent. Diluents must be free of particles and give a
background count of less than 50 particles in the measured volume. The correct
diluent for each individual instrument must be used; other diluents, even those made
by the same manufacturer, may not be interchangeable. Any laboratories using
diluents other than those recommended by the manufacturer of the instrument must
satisfy themselves that no error is being introduced.
For red cell counting in simple single-channel counters a suitable diluent requires a
pH of 7.0–7.5 and osmolality of 340 ± 10 mmol. Physiological saline (9 g/l NaCl) or
phosphate-buffered saline, which have the advantages of simplicity and ready
availability, can be used as a red cell diluent, provided that the counts are performed
immediately after dilution to avoid errors owing to sphering. Commercial solutions of
saline (for intravenous use) are usually particle-free. Other solutions may require
filtration through a 0.22- or 0.45-mm micropore filter to remove dust.
Setting Discrimination Thresholds
An accurate RBC requires that thresholds be set so that all red cells, but a minimum
of other cells, are included in the count. Some counters have a lower threshold but no
upper threshold so that white cells are included in the “RBC.” Because the WBC is
usually very low in relation to the RBC, this is not usually of practical importance;
however, an appreciable error can be introduced if the WBC is greatly elevated,
particularly if the patient is also anaemic. The setting of the lower threshold is of
considerable importance because it is necessary to ensure that microcytic red cells are
included in the count without also counting large platelets.
Current multichannel instruments, both impedance counters and counters using lightscattering technology, have thresholds that are either precalibrated by the
manufacturer or are automatically adjusted, depending on the characteristics of
individual blood samples. Single-channel impedance instruments capable of
performing a direct RBC require setting of thresholds so as to separate pulses
generated by red cells from background noise and from pulses generated by platelets.
This is done by adjusting the aperture current and the pulse amplification. A simple
method is to dilute a fresh blood sample and carry out successive counts on the
suspension, while the lower threshold control is moved incrementally from its
maximum to its minimum position. At the maximum position, the count should be
zero or close to zero, and the counts will increase as the amplitude is reduced. The
counts at each setting are plotted on arithmetic graph paper.
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Figure 9: Setting Discrimination Thresholds
Method to establish working conditions of cell counters. The correct setting of the
threshold (at arrow) is intended to exclude noise pulses without loss of the signal
pulses produced by the blood cells.
The correct threshold setting is at the left of the horizontal part of the graph before the
line begins to slope. It is important to check that the setting selected is valid for
microcytic cells. The threshold can be defined more precisely for an individual
sample by means of a pulse height analyser linked to the counting system. The lower
threshold is correctly set if beyond this point there are less than 0.5% of the counts at
the peak (mode) of the pulse size distribution curve.
2.6 Packed Cell Volume and Mean Cell Volume
Modern automated blood cell counters estimate PCV by technology that has little
connection with packing red cells by centrifugation. It is sometimes convenient to use
different terms to distinguish the manual and automated tests, and for this reason the
International Council for Standardization in Haematology has suggested that the term
“haematocrit” (Hct) rather than PCV should be used for the automated measurement.
72 | P a g e
However, it should be noted that, in the past, the terms “packed cell volume” and
“haematocrit” have been used interchangeably for the manual procedure.
With automated instruments, the derivation of the RBC, PCV, and MCV are closely
interrelated. The passage of a cell through the aperture of an impedance counter or
through the beam of light of a light-scattering instrument leads to the generation of an
electrical pulse the height of which is proportional to cell volume. The number of
pulses generated allows the RBC to be determined, as discussed earlier. Pulse height
analysis allows either the MCV or the PCV to be determined. If the average pulse
height is computed, this is indicative of the MCV, and the PCV can be derived by
multiplying the estimated MCV by the RBC. Similarly, if the pulse heights are
summated, this figure is indicative of the PCV, and the MCV can, in turn, be derived
by dividing the PCV by the RBC.
Automated instruments require calibration before the PCV or MCV can be
determined. Calibration of the PCV can be based on manual PCV determinations.
Alternatively, the MCV can be calibrated by means of the pulse heights generated by
latex beads, sdtabilize cells, or some other calibrant containing particles of known
size; however, unfixed human red cells that are biconcave and flexible will not
necessarily show the same characteristics in a cell counter as latex particles or some
other artificial calibrant. Aperture-impedance systems measure an apparent volume
that is greater than the true volume, being influenced by a “shape factor; this factor is
less than 1.1 for young, flexible red cells; is between 1.1 and 1.2 for fixed biconcave
cells; and is about 1.5 for spheres, whether they be fixed cells or latex spheres.
The MCV, and therefore the PCV, as determined by an automated counter, will vary
with certain cell characteristics other than volume. As indicated earlier, such
characteristics include shape, which in turn is partly determined by flexibility. With
impedance counters, the normal disc-shaped red cell becomes elongated into a cigar
shape as it passes through the aperture; this is caused by deformation in response to
shear force, which occurs in cells of normal flexibility. Cells with a reduced
haemoglobin concentration undergo more elongation than normal cells; this leads to a
reduced “shape factor,” a reduced pulse height in relation to the true size of the cell,
and underestimation of the MCV. Conversely, cells with abnormally rigid membranes
and cells such as spherocytes with a high haemoglobin concentration will undergo
less deformation than normal and the MCV will be overestimated. Earlier lightscattering instruments also underestimated the volume of red cells with a reduced
haemoglobin concentration because light scattering was affected by the haemoglobin
concentration. These artefacts are seen even with normal red cells of varying
haemoglobin concentration but are more apparent with red cells from patients with
defects in haemoglobin synthesis such as those from patients with iron deficiency.
Light-scattering instruments have been developed to avoid artefacts of this type. Cells
are isovolumetrically sphered so that their light-scattering characteristics are uniform
and should follow the laws of physics.
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The automated MCV and PCV are prone to certain errors that do not occur or are less
of a problem with manual methods. These include those resulting from microclots or
partial clotting of the specimen, extreme microcytosis, and the presence of
cryoglobulins or cold agglutinins; the last is a relatively common cause of factitious
elevation of the MCV because clumps of cells are sized as if they were single cells.
Because the RBC is underestimated, the PCV is less affected, although it is also
inaccurate. It is rare for warm agglutinins to cause a similar problem. Sickling may
cause a factitious increase in MCV and PCV, whereas alterations in plasma
osmolarity occurring, for example, in severe hyperglycemias also cause factitious
elevation of the MCV and PCV.
2.7 VARIATIONS IN RED CELL VOLUMES: RED CELL DISTRIBUTION
WIDTH
Automated instruments produce volume distribution histograms that allow the
presence of more than one population of cells to be appreciated. Instruments may also
assess the percentage of cells falling above and below given MCV thresholds and
“flag” the presence of an increased number of microcytes or macrocytes. Such
measurements may indicate the presence of a small but significant increase in the
percentage of either microcytes or macrocytes before there has been any change in the
MCV.
The RDW is derived from pulse height analysis and can be expressed either as the
standard deviation (in fl) or as the coefficient of variation (CV) (%) of the
measurements of the red cell volume. Current Beckman-Coulter and Bayer-Technicon
instruments express the RDW as the SD, and Sysmex instruments express it as either
the SD or the CV. The normal reference range is in the order of 12.8 ± 1.2% as CV
and 42.5 ± 3.5 fl as SD. However, widely different ranges have been reported;
therefore it is important for laboratories to determine their own reference ranges. The
RDW expressed as the CV has been found of some value in distinguishing between
iron deficiency (RDW usually increased) and thalassaemia trait (RDW usually
normal) and between megaloblastic anaemia (RDW often increased) and other causes
of macrocytosis (RDW more often normal).
2.8 TOTAL WHITE BLOOD CELL COUNT
The total WBC is determined in whole blood in which red cells have been lysed. The
lytic agent is required to destroy the red cells and reduce the red cell stroma to a
residue that causes no detectable response in the counting system without affecting
leucocytes in such a manner that the ability of the system to count them is altered.
Various manufacturers recommend specific reagents, and for multichannel
instruments that also perform an automated differential count use of the recommended
reagent is essential.
Fully automated multichannel instruments perform WBCs by impedance or lightscattering technology or both. Residual particles in a diluted blood sample are counted
after red cell lysis or, in the case of some light-scattering instruments, after the red
74 | P a g e
cells have been rendered transparent. Thresholds are set to exclude normal platelets
from the count, although giant platelets are included. Some or all of any nucleated red
cells present are usually included, so that when nucleated red cells are present the
count approximates more to the TNCC than to the WBC.
Factitiously low automated WBCs occasionally occur as a consequence of leucocyte
agglutination, prolonged sample storage, or abnormally fragile cells (e.g., in
leukaemia). Factitiously high counts are more common and usually result from failure
of lysis of red cells. With certain instruments this may occur with the cells of neonates
or be consequent on uraemia or on the presence of abnormal haemoglobin such as
haemoglobin S or haemoglobin C; high counts may also be the result of microclots,
platelet clumping, or the presence of a cryoglobulin.
2.9 AUTOMATED DIFFERENTIAL COUNT
Most automated differential counters that are now available use flow cytometry
incorporated into a full blood counter rather than being stand-alone differential
counters. Increasingly, automated blood cell counters have a differential counting
capacity, providing either a three-part or a five- to seven-part differential count.
Counts are performed on diluted whole blood in which red cells are either lysed or are
rendered transparent. A three-part differential count assigns cells to categories usually
designated:
(a) “Granulocytes” or “large cells”;
(b) “Lymphocytes” or “small cells”; and
(c) “Monocytes,” “mononuclear cells,” or “middle cells.”
In theory, the granulocyte category includes eosinophils and basophils, but in practice
it is common for an appreciable proportion of cells of these types to be excluded from
the granulocyte category and to be counted instead in the monocyte category.
Five- to seven-part differential counts classify cells as neutrophils, eosinophils,
basophils, lymphocytes, and monocytes and in an extended differential count may
also include large immature cells (composed of blasts and immature granulocytes)
and atypical lymphocytes (including small blasts).
Automated instruments performing three-part or five-to seven-part differential counts
are able to “flag” or reject counts from the majority of samples with nucleated red
cells, myelocytes, promyelocytes, blasts, and atypical lymphocytes. To a lesser extent,
instruments incorporating a three-part differential count, although not capable of
enumerating eosinophils or basophils, are able to flag a significant proportion of
samples that have an increased number of one of these cell types.
Automated differential counters using flow cytometry classify far more cells than is
possible with a manual differential count. Automated counts are consequently much
more precise than manual counts; however, with certain cell categories—specifically
monocytes and basophils—the degree of precision is sometimes less than would be
expected for the number of cells counted, indicating that such cells are not always
classified in a consistent manner. The accuracy of automated counters is less
impressive than their precision. With all types of counters, unusual cell characteristics
75 | P a g e
or ageing of a blood specimen can lead to misclassification of cells. Although the
majority of samples containing abnormal cells are “flagged,” this is not invariably so;
the presence of nucleated red cells, immature granulocytes, atypical lymphocytes, and
blasts (even occasionally quite large numbers of blasts) may not give rise to a “flag.”
However, human observers performing a 100-cell manual differential count also miss
significant abnormalities. In general, automated counts have compared favourably
with routine manual counts, especially if the instruments are assigned only two
functions—performing differential counts on normal samples and “flagging”
abnormal samples. If morphological abnormalities are flagged, microscopic
examination of a stained blood film should always be undertaken.
In the presence of a significant number of NRBC, the total count is neither a true
“WBC” nor a true “TNCC” and the absolute WBC counts calculated from the total
will necessarily be somewhat erroneous.
Instruments currently in use that count NRBC and correct the WBC for NRBC
interference include the Abbott CellDyn 4000, the Sysmex XE2100, and the
Beckman-Coulter LH750.
Differential counters based on pattern recognition in stained blood films were initially
preferred by many haematologists, but they were relatively slow, and because they
could count only a small number of cells in a reasonable time, the precision of the
automated count was no better than that of a manual count. However, with improved
computing technology and with the use of artificial neural networks, such instruments
(e.g., DiffMaster, CellVision AB, SE-223 70, and Lund, Sweden) are now capable of
providing a useful differential count on blood samples containing abnormal cells. Up
to 30 films an hour can be processed and reviewed, and abnormal cells can be
reclassified if required.
2.10 New White Cell Parameters
Many instruments are able to “flag” the presence of atypical or “variant” lymphocytes
by features such as alteration in size and in impedance or light-scattering
characteristics. Automated white cell counters can also analyse cell characteristics by
novel technologies and identify cell types by features that differ greatly from those
used when a blood film is examined visually. It is possible, for example, to identify
eosinophils by the ability of their granules to polarize light or to detect a left shift or
the presence of blasts by the reduced light scattering of the nuclei of more immature
granulocytes. There is also the potential to produce information that is not directly
analogous with that available from a manual differential count. Instruments that
incorporate a cytochemical reaction give information on enzyme activity expressed as
the mean peroxidase activity index (MPXI). An increased MPXI has been observed in
infections, in some myelodysplasias and leukaemias, in the acquired immune
deficiency syndrome (AIDS), and in megaloblastic anaemia, whereas a reduced MPXI
occurs in inherited and acquired neutrophil peroxidase deficiency. It has been
suggested that the parameters used for defining leucocyte types might also allow
detection of the presence of malaria pigment as a screening test in areas where malaria
is prevalent.
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2.11 PLATELET COUNT
Platelets can be counted in whole blood using the same techniques of electrical or
electro-optical detection as are used for counting red cells. An upper threshold is
needed to separate platelets from red cells, and a lower threshold is needed to separate
platelets from debris and electronic noise. Recirculation of red cells near the aperture
should be prevented, as pulses produced may simulate those generated by platelets.
Three techniques for setting thresholds have been used:
(a) Platelets can be counted between two fixed thresholds (e.g., between 2 and 20 fl;
(b) Pulses between fixed thresholds can be counted with subsequent fitting of a curve
and extrapolation so that platelets falling outside the fixed thresholds are included in
the computed count.
(c) Thresholds can vary automatically, depending on the characteristics of individual
blood samples, to make allowance for microcytic or fragmented red cells or for giant
platelets.
A new method for platelet counting by flow cytometry has been developed Platelets
in a blood sample are labelled fluorescently with a specific monoclonal antibody or
combination of antibodies, and by measuring the RBC :platelet ratio the platelet count
can be calculated. Suitable antibodies to platelet antigens are CD41, CD42, and
CD61. This method using CD41 and CD61 has been adopted by the International
Council for Standardization in Haematology as the reference method some
instruments now provide an automated immunological platelet count for diagnostic
use. Although these instruments can count platelets down to levels of 10 × 109/l or
less, it should be noted that precision at these levels is often poor with CVs of 22–
66% being observed and with mean counts differing appreciably between instruments.
Factitiously low automated platelet counts may be the result of giant platelets being
identified as red cells, EDTA-induced platelet clumping, and satellitism. Misleadingly
high platelet counts may be due to markedly microcytic or fragmented red cells, to
cell fragments in leukaemia, or to bacteria or fungi.
2.12 Mean Platelet Volume
The same techniques that are used to size red cells can be applied to platelets. The
calculated mean platelet volume (MPV) is very dependent on the technique of
measurement and on length and conditions of storage prior to testing the blood. When
MPV is measured by impedance technology, it has been found to vary inversely with
the platelet count in normal subjects. If this curve is extrapolated, it has been found
that data fit the extrapolated curve when thrombocytopenia is caused by peripheral
platelet destruction; however, the MPV is lower than predicted when
thrombocytopenia is caused by megaloblastic anaemia or bone marrow failure. The
MPV is generally greater than predicted in myeloproliferative disorders, but
differentiating essential thrombocythaemia from reactive thrombocytosis on this basis
has not been very successful.
Other platelet parameters that can be computed by automated counters include the
platelet distribution width (PDW), which is a measure of platelet anisocytosis, and the
77 | P a g e
“plateletcrit,” which is the product of the MPV and platelet count and, by analogy
with the haematocrit, may be seen as indicative of the volume of circulating platelets
in a unit volume of blood. The PDW has been found to be of some use in
distinguishing essential thrombocythaemia (PDW increased) from reactive
thrombocytosis (PDW normal). The plateletcrit does not appear to provide any
information of clinical value.
2.13 RETICULOCYTE COUNT
Automated reticulocyte counts have been developed by using the fact that various
dyes and fluorochromes combine with the RNA of reticulocytes. Following binding of
the dye, fluorescent cells can be enumerated using a flow cytometer. Most fully
automated blood counters now incorporate a reticulocyte counting capacity so that use
of a stand-alone reticulocyte counter is no longer necessary and use of a general
purpose flow cytometer is no longer appropriate. An international standard for this
method has been published by ICSH. The dyes used in the different systems include
auramine O (Sysmex), thiazole orange (ABX), CD4K 530 (Abbott), as well as
nonfluorescent dyes such as oxazine 750 (Bayer-Technicon) and the traditional New
methylene blue (Beckman-Coulter, Abbott).
After staining, it is necessary to separate the reticulocytes from unstained red cells,
and, because the dyes also combine with DNA of nucleated cells, these cells must also
be excluded.
The threshold for this exclusion is determined by the intensity of fluorescence and
particle sizing. Although the separation of reticulocytes from mature red cells is not
always clearcut, automated reticulocyte counts correlate well with manual reticulocyte
counts, although absolute counts may differ because automated counts are dependent
on the conditions of incubation and the method of calibrating the instrument.
Precision is much superior to that of the manual count because many more cells are
counted and the subjective element inherent in recognizing late reticulocytes is
eliminated. Potential sources of inaccuracy are the inclusion of some leucocytes and
platelets and, less often, Howell–Jolly bodies or malarial parasites in the
“reticulocyte” count.
Automated reticulocyte counts are fairly stable in blood that has been stored for 1–2
days at room temperature or up to 3–5 days at 4°C.
2.14 Immature Reticulocyte Fraction
Fully automated instruments provide a measure of the various degrees of reticulocyte
maturation because the most immature reticulocytes, produced when erythropoietin
levels are high, have more RNA and fluoresce more strongly than the mature
reticulocytes normally present in the peripheral blood. Parameters indicating
reticulocyte immaturity have potential clinical relevance. For example, an increase in
mean fluorescence intensity indicative of the presence of immature reticulocytes has
been noted as an early sign of engraftment following bone marrow transplantation.
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The characteristics of reticulocyte output in different types of anemia’s can be
especially appreciated from an output bivariate graph relating fluorescent intensity to
reticulocyte count. As described earlier,
low total count with a relatively high immature reticulocyte fraction (IRF) is
indicative of a repopulating marrow,
whereas a reticulocytopenia with low IRF is typical of severe aplastic anaemia or
renal failure
A high total count with high IRF occurs in acute haemolysis and blood loss,
whereas a low to normal total count with a high IRF occurs in dyserythropiesis and in
early response to haematinics. The appearance of reticulocytes with high
fluorescence also heralds response when severe aplastic anaemia is being treated with
immunosuppressive therapy
2.15 CALIBRATION OF AUTOMATED BLOOD CELL COUNTERS
The following methods are recommended for calibrating an automated blood cell
counter
1.
By using fresh normal blood specimens to which values have been assigned
for Hb, PCV, RBC, WBC, and platelet count by standardized reference
methods
2.
By use of a stable calibrant (either preserved blood or a substitute) to which
values appropriate for the instrument in question have been assigned by
comparison with fresh normal blood
3.
By use of a commercial calibrant with assigned values suitable for the
instrument in question
For reasons of convenience and economy, control materials are commonly used as
calibrants; but this practice is not recommended. Such materials are not sufficiently
stable to serve as calibrants and their stated values are often approximations that are
not assigned by reference methods. They are designed to give test results within a
stated range over a stated period rather than a specific result.
The procedure for assigning values to fresh blood samples and indirectly to a stable
calibrant is as follows:
1.
4 ml blood specimens are obtained from three haematologically normal
volunteers and are anticoagulated with K2 EDTA.
2.
The Hb value is assigned by using the haemiglobincyanide method and the
mean of two measurements.
3.
The PCV is assigned by the microhaematocrit method, taking the mean of
measurements in four microhaematocrit tubes.
4.
The RBC is assigned by performing counts on a single-channel apertureimpedance counter capable of performing a direct cell count; the mean of two
dilutions, each counted twice, is used.
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5.
The MCV is assigned by calculation from the RBC and PCV.
6.
The WBC is assigned by performing counts on a single-channel apertureimpedance instrument capable of performing direct cell counts; the mean of
two dilutions, each counted twice, is used.
2.16 FLAGGING OF AUTOMATED BLOOD COUNTS
“Flagging” refers to a signal that the specimen being analysed may have a significant
abnormality because one or more of the blood count variables are outside specified
limits (usually 2SD) or there is a qualitative abnormality that requires a quality
control check and/or additional investigation. This usually includes a blood film
review. Although it is theoretically desirable for every blood count to include
examination of a stained film, this is being challenged by increasing workloads
requiring time- and cost-effective rationalization, as well as by the use of automated
analyzers that report differential leucocyte counts on every specimen. Consequently,
fewer blood films are being examined microscopically. Thus, a decision of when a
blood film should be made, stained, and examined should take account of flagging
and the need to ensure analytic reliability. This includes a check of any significant
changes from a recent previous count (delta-check), as well as any specific clinical
circumstances. The following is a guide to this selection.
Blood count request: Is it a first time count or repeat count?
First time count: Is it a routine screening test or special category?
If Routine: Analyzer report for blood count alone
Film required if any flags are signalled
If Special category: Film required:
1.
Diagnosed blood disease patients
2.
Patients receiving radiotherapy and/or chemotherapy
3.
Renal disease
4.
Neonates
5.
Intensive care unit
6.
If special tests have also been requested for: infectious mononucleosis,
haemolytic anaemia, enzymopathy, abnormal haemoglobins
7.
If the clinical details on the request form indicate lymphadenopathy,
splenomegaly, jaundice or suggest the possibility of leukaemia or lymphoma
8.
Specific requests by clinician
Repeat count: Film required:
1.
Delta check positive when compared with previous record
2.
Any flag occurs in present count
3.
On each occasion for patients with known blood diseases, for neonates, and
when specifically requested by clinicians
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Module 2
Session 2: Manual techniques
Objectives
At the end of this session participants will be able to:
1. Estimate hemoglobin concentration
2. Estimate PCV or Hct
3. Estimate total WBC count
4. Estimate platelet count
5. Estimate ESR
6. Estimate reticulocyte count
7. Apply preparation of blood films on slides
8. Explain antiglobulin or coombs test
9. Determine the blood ABO and Rh group
Trainers Preparation:




Review the reading material and the session plan.
Prepare the presentation as appropriate and as recommended in the method
column of the session plan, or write the information on a flipchart or board
where all participants can see it.
Prepare copies of the reference materials/handouts and exercises.
Arrange the training room.
Methods and activities

Mini-lecture, discussion, question & answer, brain storming
Evaluation/assessment
Questions and answers, trainer’s observation
Estimated Time
255 minutes
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2.2.1. Estimate
hemoglobin
concentration
2.2.2. Estimate PCV or
Hct
Mini-lecture

The hemoglobin
concentration (Hb)
may be estimated by
measurement of its
color, by its power of
combining with
oxygen or carbon
monoxide, or by its
iron content

Reference range
Men =150 +_ 20g/l
Women =135 +_
15g/l

The haematocrit
×1000 is about
three times the
haemoglobin
expressed in g/l
In conjunction with
estimations of
haemoglobin and
red blood cell count
(RBC), it can be
used in the
calculation of red
cell indices.
Factors affecting
the test:
-Factors decerasing
microhaematocrit
- Factors increasing
microhaematocrit
Reference rage
Men =0.45+_
0.05(l/l)
Women =
0.41+_0.05(l/l)



30 minutes
Discussion
30 minutes
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2.2.3. Estimate total
WBC count
Question & answers

Visual counting of
blood cells is an
acceptable
alternative to
electronic counting
for white cell, while,
it is not
recommended for
routine red cell
counts because the
number of cells that
can be counted
within a reasonable
time in the routine
laboratory will be too
few to ensure a
sufficiently precise
result

Factors affecting
the test
Range of WBC
count in health
4.0–10.0 × 109/l
2.2.4. Estimate platelet
count

30 minutes
Question & answers
30 minutes

Manual counts are
used routinely in
under-resourced
laboratories, and
they are still needed
even in wellequipped
laboratories for blood
samples with a
significant proportion
of giant platelets

Factors affecting
the test
Range of platelet
Count
280 ± 130 × 109/l

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Mini-lecture
2.2.5 Estimate ESR
30 minutes



2.2.6. Estimate
reticulocyte count
Question & answers



2.2.7. Apply preparation
of blood films on slides
Essentially it is the
measurement after 1
hour of the
sedimentation of
red cells in diluted
blood in an openended glass tube of
30 cm length
mounted vertically
on a stand.
Range in health
- Men
- Women
- Pregnancy
Factors affecting
the test
Reticulocytes are
juvenile red cells;
they contain
remnants of the
ribosomal
ribonucleic acid
(RNA) that was
present in larger
amounts in the
cytoplasm of the
nucleated
precursors from
which they were
derived
Factors affecting
the test
Range of
Reticulocyte Count
in Health
50–100 × 109/l (0.5–2
30 minutes
Mini-lecture
84 | P a g e



Blood films should 30 minutes
be made on clean
glass slides.
Films made on
cover glasses have
negligible
advantages and are
unsuitable for
modern laboratory
practice.
Films may be
spread by hand or
by means of an
automated slide
spreader, the latter
being either a standalone instrument or
a component of an
automated blood
cell counter.
2.2.8. Explain
antiglobulin or coombs
test
Mini-lecture



The antiglobulin
test is probably the
most important test
in serology and
blood banking
DAT
IAT
25 minutes
Brain storming
2.2.9. Determine the
blood ABO and Rh group



20 minutes
Materials
Method
Discrepancies
85 | P a g e
Manual Techniques
4.3.1 Hemoglobin Estimation
Introduction
The hemoglobin concentration (Hb) may be estimated by measurement of its colour,
by its power of combining with oxygen or carbon monoxide, or by its iron content.
The procedures to be described are all colour or light-intensity matching techniques,
which also measure, to a varying extent, any methaemoglobin (Hi) or
sulphaemoglobin (SHb) that may be present.
Clinical significance
This test is done to detect if the patient is suffering from anaemia or rather high Hb
Concentration as in polycythaemia.
Principle:
To measure hemoglobin concentration (Hb), a known volume of carefully mixed whole
blood is added to a diluent which lyses red cells to produce a hemoglobin solution; lysis
occurs because of the hypotonicity of the diluent. The Hb is then determined from the
light absorbance (optical density) of the solution of hemoglobin or its derivative at a
selected wavelength.
Specimen
Venous or free-flowing capillary blood that has been anticoagulated with:
ethylenediaminetetra-acetic acid (EDTA).
Glassware and equipment
EDTA tubes, dispenser or micropipette, test tubes, spectrophotometer or colorimeter.
HAEMIGLOBINCYANIDE (CYANMETHAEMOGLOBIN) PROCEDURE
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The haemiglobincyanide (Cyanmethemoglobin) procedure is the internationally
recommended procedure for determining the haemoglobin concentration of blood. The
basis of the procedure is dilution of blood in a solution containing potassium cyanide
and potassium ferricyanide. Haemoglobin, Hi, and HbCO, but not SHb, are converted
to HiCN. The absorbance of the solution is then measured in a spectrometer at a
wavelength of 540 nm or a photoelectric colorimeter with a yellow–green filter.
Reagents:
Diluent
The original (Drabkin's) reagent had a pH of 8.6. The following modified solution,
Drapkin-type reagent, as recommended by the International Committee for
Standardization in Haematology, has a pH of 7.0–7.4. It is less likely to cause
turbidity from precipitation of plasma proteins and requires a shorter conversion time
(3–5 min) than the original Drabkin's solution, but it has the disadvantage that the
detergent causes some frothing:
Potassium ferricyanide (0.607 mmol/l)
200 mg
Potassium cyanide (0.768 mmol/l)
50 mg
Potassium dihydrogen phosphate (1.029 mmol/l) 140 mg
Nonionic detergent
1 ml
Distilled or deionized water
To 1 litre
The pH should be 7.0–7.4 and must be checked with a pH meter at least once a month.
The diluent should be clear and pale yellow in colour. When measured against water as
a blank in a spectrometer at a wavelength of 540 nm, absorbance must be zero. If stored
at room temperature in a brown borosilicate glass bottle, the solution keeps for several
months. If the ambient temperature is higher than 30°C, the solution should be stored
in the refrigerator but brought to room temperature before use. It must not be allowed
to freeze. The reagent must be discarded if it becomes turbid, if the pH is found to be
outside the 7.0–7.4 range, or if it has an absorbance other than zero at 540 nm against
water blank.
Procedure
Make a 1 in 201 dilution of blood by adding 20 μl of blood to 4 ml of diluent, or
according to the manufactures instructions Stopper the tube containing the solution and
invert it several times. Let the test sample stand at room temperature for at least 5 min
(to ensure the complete conversion of haemoglobin to haemiglobinocyanide), and then
pour it into a cuvette and read the absorbance in a spectrometer at 540 nm or in a
photoelectric colorimeter with a suitable filter, against a reagent blank. The absorbance
of the test sample must be measured within 6 hours of its initial dilution. The absorbance
of a commercially available HiCN standard (brought to room temperature if previously
stored in a refrigerator) should also be compared to a reagent blank in the same
spectrometer or photoelectric colorimeter as the patient sample. The standard should be
87 | P a g e
kept in the dark, and, to ensure that contamination is avoided, any unused solution
should be discarded at the end of the day on which the ampoule is opened.
Calculation of Haemoglobin Concentration
Preparation of Standard Graph and Standard Table
When many blood samples are to be tested, it is convenient to read the results from a
standard graph or table relating absorbance readings to haemoglobin in g/l for the
individual instrument. This graph should be prepared each time a new photometer is
put into use or when a bulb or other components are replaced. It can be prepared as
follows:
Prepare five dilutions of the HiCN reference standard (or equivalent preparation)
(brought to room temperature) with the cyanide–ferricyanide reagent according to the
Table below because the graph will be used to determine the haemoglobin
measurements, it is essential that the dilutions are performed accurately.
Dilutions of haemiglobincyanide (HiCN) reference solution for preparation of
standard graph
Haemoglobin [*] HiCN volume
Tube (%)
(ml)
Reagent volume (ml)
1
100 (full strength) 4.0 (neat)
None
2
75
3.0
1.0
3
50
2.0
2.0
4
25
1.0
3.0
5
0
None
4.0 (neat)
*
As percent of haemoglobin in reference solution.
The haemoglobin concentration of the reference preparation in each tube should be
plotted against the absorbance measurement. For example, if the label on the reference
preparation states that it contains 800 mg/l, (i.e. 0.8 g/l) and the procedure for
haemoglobin measurement uses a dilution of 1:201, the respective haemoglobin
concentrations of tubes 1–5 would be 160 g/l, 120 g/l, 80 g/l, 40 g/l, and zero.
Using linear graph paper, plot the absorbance values on the vertical axis and the
haemoglobin values on the horizontal axis. The points should fit a straight line that
passes through the origin. Providing that the standard has been correctly diluted, this
provides a check that the calibration of the photometer is linear. From the graph, it is
possible to construct a table of readings and corresponding haemoglobin values. This
is more convenient than reading values from a graph when large numbers of
measurements are made. It is important that the performance of the instrument does not
vary and that its calibration remains constant in relation to haemoglobin measurements.
To ensure this, the reference preparation should be measured at frequent intervals,
preferably with each batch of blood samples.
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Supplementary information
Inaccuracy in estimation of Hb due to:
The slow conversion of carboxyhemoglobin to methemoglobin leads to overestimation
of the Hb if the test is read at 3 minutes, since carboxyhemoglobin absorbs more light
at 540 nm than does cyanmethemoglobin. This difficulty can be overcome by
prolonging the reaction time to 30 min before reading. The difference between the 5
and 30 min readings can be used as a semiquantitative procedure for estimating the
percentage of HbCO in the blood.
Spectrophotometers and photometers are both sensitive to the effect of turbidity,
which may be caused by a high WBC, high concentrations of lipids or plasma proteins,
or non-lysed red cells. Increased turbidity causes a factitiously elevated estimate of Hb.
When the WBC is high, turbidity effects are circumvented by centrifugation or
filtration of the solution prior to reading the absorbance.
When turbidity is due to a high level of plasma protein, it can be cleared by the
addition of either potassium carbonate or a drop of 25% ammonia solution.
When turbidity is due to hyperlipidemia, a blank can be prepared from the diluent and
the patient’s plasma or the lipid can be removed by diethyl ether extraction and
centrifugation.
Calculation of Haemoglobin Concentration
Reference range
Haemoglobin
Men
150 ± 20 g/l
Women 135 ± 15 g/l
Units
g / l, mg/dl
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4.3.2 PACKED CELL VOLUME OR HAEMATOCRIT
Introduction
The haematocrit ×1000 is about three times the haemoglobin expressed in g/l. In
conjunction with estimations of haemoglobin and red blood cell count (RBC), it can be
used in the calculation of red cell indices. However, its use in under-resourced
laboratories may be limited by the need for a specialized centrifuge and a reliable supply
of capillary tubes.
Clinical significance
The packed cell volume (PCV) can be used primarily as a simple screening test for
anaemia, as a reference procedure for calibrating automated blood count systems, and
as a rough guide to the accuracy of haemoglobin measurements.
Principle
The centrifuge used for the capillary tubes provides a relative centrifugal force (g) of
about 12000 g, and 5 min centrifugation results in precipitation of RBC in the distal
end.
Specimen
Anticoagulated blood sample, because the PCV gradually increases with storage, the
test should be performed within 6 hours of collecting the blood sample, but a delay of
up to 24 hours is acceptable if the blood is kept at 4°C.
Failure to mix the blood sample adequately will produce an inaccurate result. The
degree of oxygenation of the blood also affects the result because the PCV of venous
blood is ∼2% higher than that of fully aerated blood (which has lost CO2 and taken up
O2). To ensure adequate oxygenation and sample mixing, the free air space above the
sample should be >20% of the container volume.
Glassware and equipment
Anticoagulant
K2-EDTA tube is recommended, because K3-EDTA causes shrinking of the red cells,
reducing the PCV by about 2%. Anticoagulant concentration in excess of 2.2 mg/ml
may also cause a falsely low PCV as a result of cell shrinkage.
Capillary Tubes
Variation of the bore of the tubes may cause serious errors if they are not within the
narrow limits of defined specifications that should be met by manufacturers: length 75
± 0.5 mm; internal diameter 1.07–1.25 mm, wall thickness 0.18–0.23 mm; and bore
taper not exceeding 2% of the internal diameter over the entire length of the tube.
90 | P a g e
Centrifuge
Centrifuges should be checked at intervals (at least annually) by a tachometer for speed
and by a stopwatch for timer accuracy. Efficiency of packing should also be tested by
centrifuging samples of normal and polycythaemic blood for varying times from 5 to
10 min to determine the minimum time for complete packing of the red cells.
Procedure
MICROHAEMATOCRIT PROCEDURE
The microhaematocrit procedure is carried out on blood contained in capillary tubes
75 mm in length and having an internal diameter of about 1 mm. The tubes may be
plain for use with anticoagulated blood samples or coated inside with 1 iu of heparin
for the direct collection of capillary blood. The centrifuge used for the capillary tubes
provides a centrifugal force of c 12000 g, and 5 min centrifugation results in a constant
PCV. When the PCV is greater than 0.5, it may be necessary to centrifuge for a further
5 min.
Allow blood from a well-mixed specimen, or from a free flow of blood by skin
puncture, to enter the tube by capillarity, leaving at least 15 mm unfilled. Then seal the
tube by a plastic seal (e.g. Cristaseal, Hawksley, and Lancing, Sussex). Sealing the tube
by heating is not recommended because the seals tend to be tapered and there is the
likelihood of lysis. After centrifugation for 5 min, measure the proportion of cells to the
whole column (i.e., the PCV) using a reading device.
Accuracy of Microhaematocrit
The microhaematocrit procedure has an adequate level of accuracy and precision for
clinical utility. Attention must be paid to a number of factors that may produce an
inaccurate result.
Reading
The test should be read as soon as possible after centrifugation because the red cells
begin to swell and the interface becomes progressively more indistinct. To avoid errors
in reading with the special reading device, a magnifying glass should be used. White
cells and platelets (the buffy coat) must be excluded as far as possible from the reading
of the packed red cells. If a special reading device is not available, the ratio of red cell
column to whole column can be calculated from measurements obtained by placing the
tube against arithmetic graph paper or against a ruler.
Supplementary information
Plasma Trapping
The amount of plasma trapped between red cells, especially in the lower end of the red
cell column, and red cell dehydration during centrifugation generally counterbalance
each other, and the error caused by trapped plasma is usually not more than 0.01 PCV
units. Thus, in routine practice, it is unnecessary to correct for trapped plasma, but if
91 | P a g e
the PCV is required for calibrating a blood cell analyser or for calculating blood
volume, the observed PCV should be reduced by a 2% correction factor after it has been
centrifuged for 5 min or for 10 min with polycythaemic blood. It is, however, preferable
to use the surrogate reference procedure. Plasma trapping is increased in macrocytic
anaemias, spherocytosis, thalassaemia, hypochromic anaemias, and sickle cell anaemia;
it may be as high as 20% in sickle cell anaemia if all the cells are sickled.
Factors affecting the test:
Reference range
Packed cell volume (PCV) or Haematocrit (Hct)
Men
0.45 ± 0.05 (l/l)
Women 0.41 ± 0.05 (l/l)
Units
% or L / L
4.3.3 Total White Blood Cell Count
Introduction
Visual counting of blood cells is an acceptable alternative to electronic counting for
white cell, while, it is not recommended for routine red cell counts because the number
of cells that can be counted within a reasonable time in the routine laboratory will be
too few to ensure a sufficiently precise result.
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Principle:
The principles of manual cell counts, the use of the haemocytometer counting chamber
for manually counting white cells and platelets in a known surface area.
Clinical significance
Some of the common Causes of leucocytosis:
Bacterial infections
Inflammation and tissue necrosis
Metabolic disorders (e.g. uraemia, eclampsia, acidosis, gout)
Neoplasms of all types (e.g. carcinoma, lymphoma, melanoma)
Acute haemorrhage or haemolysis
Drugs (e.g. corticosteroid therapy (inhibits margination): lithium, tetracycline)
Chronic myeloid leukaemia, myeloproliferative disease, polycythaemia vera,
myelofibrosis, essential thrombocythaemia
Treatment with myeloid growth factors (e.g. G-CSF, GM-CSF)
Infectious mononucleosis, rubella, pertussis,
Infectious hepatitis, cytomegalovirus, HIV, herpes simplex or zoster
tuberculosis, toxoplasmosis, brucellosis, syphilis
Chronic lymphoid leukaemias
Acute lymphoblastic leukaemia
Non-Hodgkin's lymphoma (some)
Thyrotoxicosis.
Some of the common Causes of leucopenia:
Selective neutropenia
Drug-induced
Systemic lupus erythematosus
Felty's syndrome
Hypersensitivity and anaphylaxis
Viral (e.g. hepatitis, influenza, HIV)
Fulminant bacterial infection (e.g. typhoid, miliary Tuberculosis)
Specimen
Venous or free-flowing capillary blood that has been anticoagulated with
ethylenediamine tetra-acetic acid (EDTA). Thorough mixing of the blood specimen
before sampling is essential for accurate test results. Ideally, tests should be performed
within 6 hours of obtaining the blood specimen because some test results are altered by
longer periods of storage. However, results that are sufficiently reliable for clinical
purposes can usually be obtained on blood stored for up to 24 hours at 4°C.
Glassware and equipment
Counting Chambers
The visibility of the rulings in the counting chamber is as important as the accuracy of
calibration, so that chambers with a “metallised” surface and Neubauer or Improved
Neubauer rulings are recommended. These have nine 1 mm × 1 mm ruled areas, which,
when covered correctly with the special thick cover glass, each contain a volume of 0.1
μl of diluted blood. Coverslips designed for mounting of microscopy preparations must
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not be used with counting chambers. The sample is introduced between the chamber
and the coverglass using a pipette or capillary tube, and the preparation is viewed using
a ×40 objective and ×6 or ×10 eyepieces. With Neubauer and Improved Neubauer
chambers, count the cells in 4 or 8 horizontal rectangles of 1 mm × 0.05 mm (80 or 160
small squares) or in 5 or 10 groups of 16 small squares, including the cells that touch
the bottom and left-hand margins of the small squares.
Figure 10: Counting chamber



Microscope
Micropipette
Test tube
Reagents:
Diluent
To make the counting of white cells easier, diluted whole blood is mixed with a fluid
to lyse the red cells and stain the white cell nuclei deep violet-black. The diluent is 2%
(20 ml/l) acetic acid coloured pale violet with gentian violet.
Procedure
Make a 1 in 20 dilution of blood by adding 0.1 ml of well-mixed blood (lack of adequate
mixing is a major source of error) to 1.9 ml of diluent in a 75 × 10 mm plastic (or glass)
tube. After sealing the tube with a lid or tightly fitting bung, mix the diluted blood in a
mechanical mixer or by hand for at least 2 min by tilting the tube to an angle of about
120 degrees combined with rotation, thus allowing the air bubble to mix the suspension.
Fill a clean dry counting chamber, with its coverglass already in position, without delay.
This is simply accomplished with the aid of a plastic Pasteur pipette or a length of stout
capillary glass tubing that has been allowed to take up the suspension by capillarity.
94 | P a g e
Take care that the counting chamber is filled in one action and that no fluid flows into
the surrounding moat.
Leave the chamber undisturbed on a bench for at least 2 min for the cells to settle, but
not much longer, because drying at the edges of the preparation initiates currents that
cause movement of the cells after they have settled. The bench must be free of
vibrations, and the chamber must not be exposed to draughts or to direct sunlight or
other sources of heat. It is important that the coverglass should be of a special thick
glass and perfectly flat, so that when laid on the counting chamber, diffraction rings are
seen. The coverglass should be of such a size that when placed correctly on the counting
chamber the central ruled areas lie in the centre of the rectangle to be filled with the cell
suspension.
Factors affecting the test:
If any of the following filling defects occur, the preparation must be discarded and the
filling procedure must be repeated using another clean dry chamber:
Overflow into moat
Chamber area incompletely filled
Air bubbles anywhere in chamber area
Any debris in chamber area
To obtain a coefficient of variation of 5%, it is necessary to count about 400 cells in
practice, it is reasonable to count 100 white cells. To minimise distribution errors, count
the cells in the entire ruled area (i.e., 9 × 0.1 μl areas in an Improved Neubauer counting
chamber).
Counting in only one or two fields results in a wide variance that is reduced as more
cells are counted. However, high precision is achieved only when thousands of cells are
counted, which is only possible with automated cell counters.
Calculation
WBC/I
Thus, if N cells are counted in 0.1 μl, then the WBC/l is as follows:
(e.g., if 115 cells are counted, the WBC is 115 × 200 × 106/l = 23 × 109/l)
Range of White Blood Cell Count in Health
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White blood cell count
4.0–10.0 × 109/l
Units
× 109/l
4.3.4 PLATELET COUNT
Introduction
Manual counts are used routinely in under-resourced laboratories, and they are still needed even
in well-equipped laboratories for blood samples with a significant proportion of giant platelets.
However, for all other samples, automated full blood counters produce platelet counts with a
precision that is much superior to that of manual platelet counts.
Principle:
Manual platelet counts are performed by visual examination of diluted, lysed whole blood
using a Neubauer or Improved Neubauer counting chamber as for total white cell counts.
Clinical significance
This test is used for detection of thrombocytosis or thrombocytopenia which occurs in many
diseases.
Specimen
Platelet counts are best performed on ethylenediaminetetra-acetic acid (EDTA)-anticoagulated
blood that has been obtained by clean venipuncture. They can also be carried out on blood
obtained by skin prick, but the results are less satisfactory than those on venous blood. Skinprick platelet counts are significantly lower than counts on venous blood and less constant; a
variable number of platelets are probably lost at the site of the skin puncture.
Glassware and equipment
Counting Chambers
The visibility of the rulings in the counting chamber is as important as the accuracy of
calibration, so that chambers with a “metallised” surface and Neubauer or Improved Neubauer
rulings are recommended. These have nine 1 mm × 1 mm ruled areas, which, when covered
correctly with the special thick cover glass, each contain a volume of 0.1 μl of diluted blood.
Coverslips designed for mounting of microscopy preparations must not be used with counting
chambers. The sample is introduced between the chamber and the coverglass using a pipette or
capillary tube, and the preparation is viewed using a ×40 objective and ×6 or ×10 eyepieces.
With Neubauer and Improved Neubauer chambers, count the cells in 4 or 8 horizontal
rectangles of 1 mm × 0.05 mm (80 or 160 small squares) or in 5 or 10 groups of 16 small
squares, including the cells that touch the bottom and left-hand margins of the small squares.
96 | P a g e
Figure11: Counting Chamber
 Microscope
 Micropipette
 Test tube
Reagents:
1% ammonium oxalate, which lyses red cells, as a diluent produces a higher and more accurate
count than use of formal citrate, which leaves red cells intact.
Procedure
The diluent consists of 1% aqueous ammonium oxalate in which the red cells are lysed. This
procedure is recommended in preference to that using formal-citrate as diluent, which leaves
the red cells intact and is more likely to give incorrect results, when the platelet count is low.
Before diluting the blood sample, examine it carefully for the presence of blood clots. If these
are present, a fresh specimen should be requested because clots will cause the platelet count to
be artificially low. Make a 1 in 20 dilution of well-mixed blood in the diluent by adding 0.1 ml
of blood to 1.9 ml of ammonium oxalate diluent (10 g/l). Not more than 500 ml of diluent
should be made at a time, using carefully clean glassware and fresh glass-distilled or deionised
water. If possible, the solution should be filtered through a micropore filter (0.22 μm) and kept
at 4°C. For use, a small part of the stock is refiltered and dispensed in 1.9 ml volumes in 75 ×
12 mm tubes.
Mix the suspension on a mechanical mixer for 10–15 min. Fill a Neubauer counting chamber
with the suspension, using a stout glass capillary or Pasteur pipette. Place the counting chamber
in a moist Petri dish and leave untouched for at least 20 min to give time for the platelets to
settle.
Examine the preparation with the ×40 objective and ×6 or ×10 eyepieces. The platelets appear
under ordinary illumination as small (but not minute) highly refractile particles if viewed with
97 | P a g e
the condenser racked down; they are usually well-separated, and clumps are rare if the blood
sample has been skillfully collected. To avoid introducing into the chamber dirt particles, which
might be mistaken for platelets, all equipment must be scrupulously clean. Platelets are more
easily seen with the phase-contrast microscope. A special, thin-bottomed (1 μm) counting
chamber is best for optimal phase-contrast effect. The number of platelets in one or more areas
of 1 mm2 should be counted. The total number of platelets counted should always exceed 200
to ensure a coefficient of variation of 8–10%.
Calculation
Thus, if N is the number of platelets counted in an area of 1 mm2 (0.1 μl in volume), the
number of platelets per litre of blood is:
Range of platelet Count
Platelet count 280 ± 130 × 109/l
Factors affecting the test:
The errors associated with manual cell counts are technical and inherent.
Technical errors can be minimized by avoiding the following:

Poor technique in obtaining the blood specimen

Insufficient mixing of the blood specimen

Inaccurate pipetting and the use of badly calibrated pipettes or counting
chambers

Inadequate mixing of the cell suspension

Faulty filling of the counting chamber

Careless counting of cells within the chamber
Units
× 109/l
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4.3.5 Erythrocyte Sedimentation Rate
Introduction
Inflammatory response to tissue injury (the acute-phase response) includes alteration in
serum protein concentration, especially increases in fibrinogen, haptoglobin,
caeruloplasmin, immunoglobulins (Ig), and C-reactive protein (CRP), and decrease in
albumin. The changes occur in acute infection, during active phases of chronic
inflammation, with malignancy, in acute tissue damage (e.g., following acute
myocardial infarction), or with physical injury.
Principle:
Essentially it is the measurement after 1 hour of the sedimentation of red cells in diluted
blood in an open-ended glass tube of 30 cm length mounted vertically on a stand.
Clinical significance
Measurement of the acute-phase response is a helpful indicator of the presence and
extent of inflammation or tissue damage and response to treatment. The usual tests are
estimation of CRP and measurement of the erythrocyte sedimentation rate (ESR);
Specimen
Either collect venous blood in ethylenediaminetetra-acetic acid (EDTA) and dilute a
sample accurately in the proportion of 1 volume of citrate to 4 volumes of blood,
or collect the blood directly into the citrate solution.
Glassware and equipment
The recommended tube is a straight glass or rigid transparent plastic tube 30 cm in
length and not less than 2.55 mm in diameter. The bore must be uniform to within 5%
throughout. A scale graduated in mm extends over the lower 20 cm. The tube must be
clean and dry.
Specially made racks with adjustable levelling screws are available for holding the
sedimentation tubes firmly in an exactly vertical position. The rack must be constructed
so that there will be no leakage of the blood from the tube.
Reagents:
Diluent prepare a solution of 109 mmol/l trisodium citrate (32 g/l Na3Ca6H5O7.2H2O).
Filter through a micropore filter (0.22 mm) into a sterile bottle. It can be stored for
several months at 4°C but must be discarded if it becomes turbid through the growth of
moulds.
Procedure
Conventional Westergren Procedure
The test should then be carried out on the diluted sample within 4 h of collecting the
blood, although a delay of up to 6 h is permissible provided that the blood is kept at
4°C. EDTA blood can be used within 24 h if the specimen is kept at 4°C, provided that
1 volume of 109 mmol/l (32 g/l) trisodium citrate is added to 4 volumes of blood
immediately before the test is performed.
99 | P a g e
Mix the blood sample thoroughly and then draw it up into the Westergren tube to the
200 mm mark by means of a teat or a mechanical device; mouth suction should never
be used. Place the tube exactly vertical and leave undisturbed for exactly 60 min, free
from vibrations and draughts and not exposed to direct sunlight.
Calculation
Read to the nearest 1 mm the height of the clear plasma above the upper limit of the
column of sedimenting cells. The result is expressed as ESR = X mm in 1 h. A poor
delineation of the upper layer of red cells may sometimes occur, especially when there
is a high reticulocyte count.
Range in Health
There is a progressive increase with age, but older than 70 years it is difficult to define
a strictly healthy population for determining normal values.
Erythrocyte sedimentation rate ranges in health
Age (years) 95% Upper limit (mm in 1h)
Men
17–50
10
51–60
12
61–70
14
>70
about 30
Women
17–50
12
51–60
19
61–70
20
>70
about 35
Pregnancy
First half
48 (62 if anaemic)
Second half 70 (95 if anaemic)
In the newborn, the ESR is usually low. In childhood and adolescence, it is the same as
for normal men with no differences between boys and girls. It is increased in pregnancy,
especially so in the later stages, and independent of anaemia.
Factors affecting the test:
Modified Procedures
Length of Tube
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The overall length of the tube is not a critical dimension for the test provided that it fits
firmly in an appropriate holding device. The tube must, however, be long enough to
ensure that packing of the cells does not start before the test has been completed.
Temp: should be between 18-25 C
Hemolysis: affect the rate of sedimentation
Plastic Glass Tubes
A number of plastic materials (e.g., polypropylene and polycarbonate) are
recommended as substitutes for glass in Westergren tubes. Nevertheless, not all plastics
have similar properties, and it must be demonstrated that the ESR with the chosen tubes
is reproducible and not affected by the plastic.
Disposable Glass Tubes
Disposable glass tubes should be supplied clean and dry and ready for use. It is
necessary to show that neither the tube material nor the manufacturer's cleaning process
affect the ESR.
Capillary Procedure
Short tubes of narrower bore than in the standard tube are available mainly for tests on
infants. These are, however, no longer in general use, and it is necessary to establish
normal ranges or a correction factor to convert results to an approximation of ESR by
the Westergren procedure.
Time
Sedimentation is measured after aggregation has occurred and before the cells start to
pack usually at 18-24 min. From the rate during this time period the sedimentation that
would have occurred at 60 min is derived and converted to the conventional ESR
equivalent by an algorithm.
Sloping Tube
Red cells sediment more quickly when streaming down the wall of a sloped tube. This
phenomenon has been incorporated into automated systems in which the end-point is
read after 20 min with the tube held at an angle of 18 degrees from the vertical. This
has been shown to give results comparable to the conventional procedure.
Anticoagulant
EDTA blood can be used without citrate dilution, at least if packed cell volume (PCV)
is below 0.36 (haemoglobin < 110 g/l); less precise results are obtained when the PCV
is higher. The readings from undiluted samples must then be adjusted as for the
standardised procedure (see below).
Because of the biohazard risk of blood contamination inherent in using open-ended
tubes, it is now recommended that, where possible, a closed system be used in routine
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practice. Manual procedures are available that avoid transfer of the blood into the
sedimentation tube. Automated closed systems use either blood collected in special
evacuated tubes containing citrate or EDTA blood. A sample is taken up through a
pierceable cap and then automatically diluted in the system if this is required. Some
systems use sloping tubes at an angle of 18 degrees to obtain results rapidly, and one
model of the Ves-matic (Diesse) also incorporates centrifugation.
Whenever a different procedure or tube is planned, a preliminary test should be carried
out to check precision and to compare results with those obtained by the standardised
procedure described in the following section.
Units
Mm / hour
4.3.6 RETICULOCYTE COUNT
Introduction
Reticulocytes are juvenile red cells; they contain remnants of the ribosomal ribonucleic
acid (RNA) that was present in larger amounts in the cytoplasm of the nucleated
precursors from which they were derived. Ribosomes have the property of reacting with
certain basic dyes such as azure B, brilliant cresyl blue, or New methylene blue (see
below) to form a blue or purple precipitate of granules or filaments.
Principle
On exposure of unfixed cells to certain dyes that contains only basic component of the
stain, such as brilliant cresyl blue or new methylene blue, the ribosomes are precipitated
and stained by the dye, to appear as a reticular network. With these stains red cells stain
a pale greenish-blue while the reticulum stains bluish-purple.
Clinical significance
The number of reticulocytes in the peripheral blood is a fairly accurate reflection of
erythropoietic activity, assuming that the reticulocytes are released normally from the
bone marrow and that they remain in circulation for the normal time period.
Specimen
Venous blood that has been anticoagulated with ethylenediaminetetra-acetic acid
(EDTA). Thorough mixing of the blood specimen before sampling is essential for
accurate test results. Ideally, tests should be performed within 6 hours of obtaining the
blood specimen because some test results are altered by longer periods of storage.
However, results that are sufficiently reliable for clinical purposes can usually be
obtained on blood stored for up to 24 hours at 4°C.
Glassware and equipment
EDTA tubes, dispenser or micropipette, test tubes, slides, Microscope.
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Reagents
Staining Solution
Dissolve 1.0 g of New methylene blue or azure B in 100 ml of iso-osmotic phosphate
buffer pH 6.5.
Procedure
Deliver 2 or 3 drops of the dye solution into a 75- × 10-mm plastic tube by means of a
plastic Pasteur pipette. Add 2–4 volumes of the patient's EDTA-anticoagulated blood
to the dye solution and mix. Keep the mixture at 37°C for 15–20 min. Resuspend the
red cells by gentle mixing, and make films on glass slides in the usual way. When dry,
examine the films without fixing or counterstaining.
The exact volume of blood to be added to the dye solution for optimal staining depends
on the RBC. A larger proportion of anaemic blood, and a smaller proportion of
polycythaemic blood, should be added than of normal blood. In a successful
preparation, the reticulofilamentous material should be stained deep blue and the
nonreticulated cells should be stained diffuse shades of pale greenish blue. Films should
not be counterstained. The reticulofilamentous material is not better defined after
counterstaining, and precipitated stain overlying cells may cause confusion. Moreover,
Heinz bodies will not be visible in fixed and counterstained preparations. If the stained
preparation is examined under phase contrast, both the mature red cells and
reticulocytes are well defined. By this technique, late reticulocytes characterized by the
presence of remnants of filaments or threads are readily distinguished from cells
containing inclusion bodies. Satisfactory counts may be made on blood that has been
allowed to stand (unstained) for as long as 24 hours, although the count will tend to
decrease after 6–8 hours unless the blood is kept at 4°C.
Counting Reticulocytes
An area of film should be chosen for the count where the cells are undistorted and where
the staining is good. A common fault is to make the film too thin; however, the cells
should not overlap. To count the cells, use the ×100 oil-immersion objective and, if
possible, eyepieces provided with an adjustable diaphragm. If eyepieces with an
adjustable diaphragm are not available, a paper or cardboard diaphragm, in the centre
of which has been cut a small square with sides about 4 mm in length, can be inserted
into an eyepiece and used as a less convenient substitute.
The counting procedure should be appropriate to the number of reticulocytes present.
Very large numbers of cells have to be surveyed if a reasonably accurate count is to be
obtained when only small numbers of reticulocytes are present. When the count is less
than 10%, a convenient procedure is to survey successive fields until at least 100
reticulocytes have been counted and to count the total red cells in at least 10 fields to
determine the average number of red cells per field.
Calculation
Number of reticulocytes in n fields = x
103 | P a g e
Average number of red cells per field =
y
Total number of red cells in n fields = n
×y
Reticulocyte percentage = [x ÷ (n × y)] ×
100%
Absolute reticulocyte count = % × RBC
Thus, when the reticulocyte percentage is 3.3 and the RBC is 5 × 1012/l, the absolute
reticulocyte count per litre is as follows: [3.3/100] × 5 × 1012 = 165 × 109
It is essential that the reticulocyte preparation be well spread to ensure an even
distribution of cells in successive fields.
When the reticulocyte count exceeds 10%, only a relatively small number of cells will
have to be surveyed to obtain a standard error of 10%.
Factors affecting the test:
It is essential that the reticulocyte preparation be well spread and well stained. Other
important factors that affect the accuracy of the count are the visual acuity and patience
of the observer and the quality and resolving power of the microscope. The most
accurate counts are carried out by a conscientious observer who has no knowledge of
the supposed reticulocyte level, thus eliminating the effect of conscious or unconscious
bias.
Range of Reticulocyte Count in Health
The range of reticulocyte in adults and children is 50–100 × 109/l (0.5–2.5%). In infants
(full term, cord blood) it is 2–5%.
Carry out a rough platelet count to determine the approximate concentration, and add
an appropriate amount of the suspension to preserved blood. Mix well for 20 min and,
with continuous mixing, dispense into sterile containers. Cap and seal. At 4°C, the
preparation should have a shelf life of 3-4 months. Before use, resuspend by thorough
shaking by hand, followed by mechanical mixing for approximately 15 min.
4.3.7 PREPARATION OF BLOOD FILMS ON SLIDES
Introduction
Blood films should be made on clean glass slides. Films made on cover glasses have
negligible advantages and are unsuitable for modern laboratory practice. Films may be
spread by hand or by means of an automated slide spreader, the latter being either a
stand-alone instrument or a component of an automated blood cell counter.
Clinical significance
104 | P a g e
It’s used for diagnosis of various blood diseases.
Principle
Although we can judge the proportions of red cells and white cells in a tube of
sedimented blood, we get far more information if the blood is carefully mixed and a
thin layer is spread on a glass slide to form a blood film. The blood cells are then
preserved by exposure to the alcohol methanol, a process known as fixation. The fixed
film of blood is stained with a mixture of several dyes so that the individual cells can
be recognized when they are examined with a microscope.
Specimen
Blood films can be prepared from fresh blood with no anticoagulant added or from
ethylenediaminetetra-acetic acid (EDTA)-anticoagulated blood. Heparinized blood
should not generally be used because its staining characteristics differ from those of
EDTA-anticoagulated blood. Good films can be made in the following manner, using
clean slides, if necessary wiped free from dust immediately before use. Slides should
measure 75 × 25 mm and approximately 1 mm thick; ideally, they should be frosted at
one end to facilitate labelling, but these are more expensive.
Glassware and equipment
EDTA tubes, dispenser or micropipette, spreader, slides, Microscope.
Reagents
Leishman's Stain
Weigh out 0.2 g of the powdered dye, and transfer it to a conical flask of 200–250 ml
capacity. Add 100 ml of methanol and warm the mixture to 50°C for 15 min,
occasionally shaking it. Allow the flask to cool and filter. It is then ready for use, but it
will improve on standing.
Buffered Water
Make up 50 ml of 66 mmol/l Sörensen's phosphate buffer of the required pH to 1 litre
with water at a pH of 6.8. An alternative buffer may be prepared from buffer tablets,
which are available commercially. Solutions of the required pH are obtained by
dissolving the tablets in water.
Manual Procedure
Spreading Blood Films
First, make a spreader from a glass slide that has a smooth end. Using a glass cutter,
break off one corner of the slide, leaving a width of about 18 mm as the spreader. A
spreader can be used repeatedly unless the edge becomes chipped, but it must be
thoroughly washed and dried between films.
105 | P a g e
Place a small drop of blood in the centre line of a slide about 1 cm from one end. Then,
without delay, place a spreader in front of the drop at an angle of about 30 degrees to
the slide and move it back to make contact with the drop. The drop should spread out
quickly along the line of contact. With a steady movement of the hand, spread the drop
of blood along the slide. The spreader must not be lifted off until the last trace of blood
has been spread out; with a correctly sized drop, the film should be about 3 cm in length.
It is important that the film of blood finishes at least 1 cm before the end of the slide.
The thickness of the film can be regulated by varying the pressure and speed of
spreading and by changing the angle at which the spreader is held. With anaemic blood,
the correct thickness is achieved by using a wider angle, and, conversely, with
polycythaemic blood, the angle should be narrower. The ideal thickness is such that on
microscopy there is some overlap of red cells throughout much of the film's length. The
leucocytes should be easily recognizable throughout most of the film. With poorly made
films the leucocytes will be unevenly distributed, with monocytes and other large
leucocytes being pushed to the end and the sides of the spread. An irregular streaky film
will occur if the slide is greasy, and dust on the surface will cause patchy spots.
The films should be allowed to dry in the air. In humid conditions the films may be
exposed to a current of warm air (e.g., from a hairdryer), but this should be in a
microbiological safety hood.
Labeling Blood Films
The film should be labelled immediately after spreading. Write either a laboratory
reference number or the name of the patient and the date in pencil on the frosted end of
the slide or on the film itself (writing on the thickest part, which is least suitable for
microscopic examination). A label written in pencil will not be removed by staining. A
paper label should be affixed to the slide later. If blood films are to be stored for future
reference, apply the paper label in such a manner that it is easily read when the slides
are filed.
Fixing Blood Films
Dry the films in the air. Leave for 3 min in the fixative this must be done without delay
and the films should never be left unfixed for more than a few hours. If films are sent
to the laboratory by post, it is preferable that, when possible, they are thoroughly dried
and fixed before dispatch.
STAINING BLOOD FILMS
Romanowsky stains are used universally for routine staining of blood films, and
satisfactory results can be obtained.
Air dries the film and flood the slide with the stain. After 2 min, add double the volume
of water and stain the film for 5–7 min. Then wash it in a stream of buffered water until
it has acquired a pinkish tinge (up to 2 min). After the back of the slide has been wiped
clean, set it up right to dry.
106 | P a g e
When several batches of films are being stained in succession, the staining solution
should be renewed at intervals (e.g., after each 50 slides). Loss of staining power is
usually the result of precipitation of the eosin Y, and this will result in the nuclei
staining blue instead of purple.
Carry out a rough platelet count to determine the approximate concentration, and add
an appropriate amount of the suspension to preserved blood . Mix well for 20 min
and, with continuous mixing, dispense into sterile containers. Cap and seal. At 4°C,
the preparation should have a shelf life of 3-4 months. Before use, resuspend by
thorough shaking by hand, followed by mechanical mixing for approximately 15 min.
4.3.11 Antiglobulin or Coombs' Test
The antiglobulin test is probably the most important test in serology and blood banking. The
antiglobulin test (Coombs' test) was introduced by Coombs, Mourant, and Race in 1945 as a
method for detecting “incomplete” Rh antibodies (i.e., IgG antibodies capable of sensitizing red
cells but incapable of causing agglutination of the same cells suspended in saline) as opposed to
“complete” IgM antibodies, which do agglutinate saline-suspended red cells. Coombs' test is
divided into direct and indirect antiglobulin tests:

In the direct antiglobulin test (DAT), the patient's cells, after careful washing, are tested for
sensitization that has occurred in vivo.
 In the indirect antiglobulin test (IAT), normal red cells are incubated with a serum suspected
of containing an antibody and subsequently tested, after washing, for in vitro–bound antibody.
The DAT is used to demonstrate in vivo attachment of antibodies to red cells, as in autoimmune
hemolytic anemia, alloimmune HDN, and alloimmune hemolysis following an incompatible
transfusion.
The IAT has wide application in blood transfusion serology, including antibody screening and
identification and cross-matching.
DAT:
A spin tube technique is recommended for the routine antiglobulin test.
The test should be carried out in glass tubes. Plastic tubes are not recommended because they may
adsorb IgG, which could neutralize anti-IgG of the antiglobulin reagent. Use a blood sample in
EDTA anticoagulant.
If a cold-reacting autoantibody is present, the patient's red cells should be washed three times in
a large volume of saline warmed to 37°C to wash off cold antibodies and obtain a smooth
suspension of cells.
There is no risk of washing off adsorbed complement components. However, the washing process
should be accomplished as quickly as possible and the test should be set up immediately afterward
because bound warm antibody occasionally elutes off the cells when they are washed and falsenegative results may be obtained. “Saline” refers to 9 g/l NaCl buffered to pH 7.0.
DAT Method:Make 5% suspension of red cells that have been washed three times in saline. If
failed to wash the red cells properly then the antihuman globulin antisera may be neutralized by
immunoglobulins or complement in the surrounding serum or plasma and cause a false negative
result.
107 | P a g e
Add 1 drop of the cell suspension to 2 drops of anti-human globulin reagent. Centrifuge for 1
minute at 150g.
Examine for agglutination after gently re-suspending the button of cells. A concave mirror and
good light help in macroscopic readings. If the result appears to be negative, confirm this
microscopically.
108 | P a g e
Figure 12: Coombs Test
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4.3.12 ABO & Rh Group
The test used to determine the blood ABO and Rh group
Materials
1- Slide
2- Anti-A Monoclonal sera
3- Anti-B Monoclonal sera
4- Anti-D Monoclonal sera
5- EDTA blood
Method
123456-
Add one drop of each colored anti-sera on the slid
To each anti-sera add one drop of EDTA blood
Mix the blood with anti-sera
Rotating the slide by hand many times read the result within 2-5 mintes
Record the result
If Rh negative, test it by tube method:
A. Add one volume of antiD sera to test tube.
B. Add one volumeof 2% red cells suspension.
C. mix suspension by traping the tube.
D. leave the suspension undisturbed for 15 min and read result.
Discrepancies
A- False Positive
1- Rouleaux formation
2- Auto-immune hemolytic anemia
B- False Negative
Occurred when the anti-sera is impotent or expired
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Module three: Serology
Module Objectives:
At the end of this module the participant will be able to:
1. Apply serological tests
Modules Sessions


Session 1: RPR Test, TPHA Test, Widel Test, and Rose Bengal Test.
Session 2: CRP Test, ASO latex slide agglutination Test, RF, Toxoplasmosis Test,
Infectious mononucleosis Test, Rota Virus Latex Test, and Pregnant Test
Evaluation/ Assessment
Questions and answers, participants’ summaries, trainer’s evaluation
Estimated Training Time
5 hours
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Module 3
Session 1: RPR Test, TPHA Test, Widel Test, and Rose Bengal Test
Objectives
At the end of this session participants will be able to:
1. Understand RPR Test
2. Understand TPHA Test
3. Understand Widel Test
4. Understand Rose Bengal Test
Trainers Preparation:




Review the reading material and the session plan.
Prepare the presentation as appropriate and as recommended in the method
column of the session plan, or write the information on a flipchart or board where
all participants can see it.
Prepare copies of the reference materials/handouts and exercises.
Arrange the training room.
Methods and activities

Mini-lecture, question & answers, discussion-lecture
Evaluation/assessment
Questions and answers, trainer’s observation
Estimated Time
115 minutes
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Session plan -1-
Objectives
3.1.1. Understand RPR
Test
3.1.2. Understand TPHA
Test
3.1.3. Understand Widel
Test
3.1.4. . Understand Rose
Bengal Test for brucellosis
Content
Methodology





Principle
Material
Procedure
Interpretation
Limitation
Mini-lecture




Principle
Material
Procedure
Interpretation of
results
Question & answers



Principle
Material
Procedure


Introduction
Most species of
pathogenic for
human
Principles
Materials
Procedure
Limitation of the
procedure




25 minutes
30 minutes
Question & answers
30 minutes
Discussion-lecture
30 minutes
113 | P a g e
5. Serology
5.1Rapid Plasma Reagin (RPR) TEST
Principle
Syphilis is a chronic venereal infection caused by the spirochaete micro-organism
Treponema pallidum. As the organism cannot be cultured on artificial media, diagnosis of
syphilis depends on the detection of specific antibodies by serological tests in
combination with clinical findings.
The RPR test is “non-treponemal” in that the antibodies detected are not specific for T.
pallidum, although their presence in patient’s serum or plasma is strongly associated with
infection by the organism. This test measures antibody (IgG and IgM) produced in
response to lipoidal material released from damaged host cells as well as to lipoproteinlike material released from the spirochaetes. These antibodies tend to disappear after
successful treatment of the infection.
The RPR test consists of modified VDRL antigen containing microparticulate carbon,
which aggregates in the presence of reagin type antibodies in serum or plasma, indicating
a positive result. The aggregation can be read macroscopically. Non-reactive samples
typically appear as a smooth non-aggregated pattern, which may form buttons in the
centre of the test area .
Material
1. Carbon antigen reagent
2. Positive Control
3. Negative Control
4. Disposable Test Slides
5. Disposable Pipette Stirrers (pipstirs)
6. Dispenser
7. Needle
8. Specimen collection container
9. Timer
10. Automatic card rotator (100 rpm)
11. (0.85%) physiological saline (semi quantitative test only).
12. Test tubes for sample dilution (semi quantitative test only).
114 | P a g e
PROCEDURE
a.
1.
2.
3.
4.
5.
6.
7.
8.
9.
b.
1.
2.
3.
4.
5.
6.
Qualitative Test
Allow all reagents, controls and samples to reach room temperature before use.
Draw the sample into the pipstir taking care not to transfer any cellular elements.
Hold the pipstir over a test card circle and allow one drop (50µl) of specimen to
fall onto the test card. It is important to hold the dispenser in a vertical position
whilst dispensing the sample.
Spread the specimen evenly over the entire test circle using the mixing (broad)
end of the pipstir.
Shake the vial of carbon antigen reagent to ensure even mixing.
Attach the needle to the dispenser. Withdraw sufficient carbon antigen for the
number of tests being performed.
Keeping the dispenser in a vertical position allow one drop to fall onto the
specimen. Do not mix. Return unused antigen from the dispenser to the glass vial.
Rotate the tile for 8 minutes preferably on an automatic rotator at 100 rpm.
Read and interpret results macroscopically in good light.
Semi Quantitative Test
Prepare doubling dilutions of the sample from the undiluted specimen to 1:32
using physiological saline. Mix well.
Using the pipstirs , place one drop (50µl) of each dilution onto a separate test card
circle.
Using the mixing (broad) end of the pipstir spread each dilution evenly over the
test circle, starting at the highest dilution (1:32), proceeding to the lowest (1:2).
Continue as from 5 in the Qualitative Test.
After 8 minutes rotation, read the test and note the last circle in the dilution series
that has a positive result.
If the highest dilution tested (1:32) is reactive, proceed with a further dilution
series by preparing doubling dilutions of the sample from 1:32 to 1:512 using
physiological saline. Mix well and continue as from 2 in the semi quantitative test.
INTERPRETATION
Quality Control
It is recommended that the kit positive and negative controls are run with each batch of
test specimens. For the assay to be valid the positive control provided should give a
strong positive pattern and the negative control provided should give a clearly negative
result.
Results
115 | P a g e
Qualitative Test
Positive result
Reactive (positive) samples display characteristic agglutination ranging from slight
(weak-reactive) to intense (reactive). A strong positive reaction is seen as large
aggregates in the centre of the test circle. Weakly positive reactions are seen as small
aggregates around the edge of the test circle.
Negative result
Negative results show no aggregates. The carbon antigen either remains in a smooth
suspension or forms a distinct button.
Figure 13: Appearance of Positive and Negative RPR Results
Semi Quantitative Test
Results may be graded from strong to non-reactive and the titre expressed as the
reciprocal of the last dilution showing a positive reaction.
1. Strong Reactive (SR): Large clumps of carbon particles with a clear background.
2. Reactive (R): Large clumps of carbon particles, more dispersed than strong reactive.
3. Weak Reactive (WR): Small clumps of carbon particles with light grey background.
4. Trace Reactive (TR): Slight clumping of carbon particles, typically seen as a button of
aggregates in the centre of the test circle or dispersed around the edge of the test circle.
5. Non-Reactive (NR): A smooth grey pattern or a button of non-aggregated carbon
particles in the centre of the test circle.
116 | P a g e
Reactive samples should be recorded as antibody positive and must be subjected to
further tests to determine the presence or absence of specific anti-Treponemal antibody.
Limitations



As with all reagin tests the RPR test may give a small proportion of false positive
results. Diseases such as infectious mononucleosis, leprosy, lupus erythematous,
vaccinia and viral pneumonia can cause such reactions.
Reactive RPR test specimens should be tested with further serological tests (i.e.
TPHA and FTA-abs) since, as with any serological procedure, the diagnosis
should not be made on a single reactive result.
As with other serological tests, the RPR test cannot distinguish between syphilis
and other pathogenic Treponemal infections e.g. Yaws.
5.2 TPHA(Treponema pallidum Haemagglutination Assay ) TEST
Principle
Serological screening tests for syphilis using cardiolipin and lecithin as antigens are
simple to perform but biological false positive reactions occur frequently because the
tests use non-treponemal antigens.
TPHA reageants are used to detect human serum antibody to T.pallidum by means of an
indirect haemagglutination (IHA) method. Preserved avian erythrocytes are coated with
antigenic components of pathogenic T.pallidum . These Test Cells agglutinate in the
presence of specific antibodies to T.pallidum, and show characteristic patterns in
microtitration plates.
Any non-specific reactions occurring are detected using the Control Cells, which are
avian erythrocytes not coated with T.pallidum antigens. Non-specific reactions may also
be absorbed out using these Control Cells.
Antibodies to non-pathogenic treponemes are absorbed by an extract of Reiter’s
treponemes, included in the cell suspension. Test results are obtained in 45-60 minutes
and the cell agglutination patterns are both easily read and long lasting.
TPHA test has been shown to be a convenient and specific test for the diagnosis of
treponemal infection, having a specificity and a sensitivity comparable to that of the
FTA-ABS test (Fluorescent treponemal antibody absorption Test) . It requires minimum
laboratory equipment and is very simple to perform.
Material
1. Test cells; preserved avian erythrocytes sensitised with T.pallidum antigen.
2. Control cells;preserved avian erythrocyte.
3. Diluent.
117 | P a g e
4. Positive control serum; (prediluted 1:20)This will give an equivalent titre of
1/640:/2560 in the quantitative test.
5. Non-reactive control serum; (prediluted 1:20).
6. Accurate pipettes for delivering 10,25,75 and 190 microlitres.
7. U-Well microtitration plates.
Procedure
Qualitative method
Each sample requires 3 wells of a microtitration plate.
1.
2.
3.
4.
Add 190µl of diluent to Well 1.
Add 10µl serum to Well 1.
Using a micropipette, mix contents of Well 1 and transfer 25µl to Wells 2 and3.
Ensure that the Test and Control Cells are thoroughly resuspended. Add 75µl of
control cells to Well 2. Add 75µl of Test Cells to Well 3.
5. Tap the plate gently to mix the contents thoroughly.
6. Incubate 45-60 minutes at room temperature. ( Keep the plate away from heat,
direct sunlight and any source of vibration).
7. Read results. Results are stable for 24hrs if the plate is covered and the above
precautions are observed.
Quantitative test
Each sample requires 8 Wells of a microtitration plate. Labelled A through to H.
1. Add 25µl of diluent to Wells B to H inclusive.
2. Transfer 25µl of 1:20 serum dilution from screening test to Wells A and B.
3. Take 25µl of diluted serum from Well B and serially dilute from Wells B to H
inclusive in 25µl aliquots, discarding 25µl of diluted serum from Well H.
4. Ensure that the Test Cells are thoroughly resuspended. Add 75µl of Test cells to
wells A to H inclusive. This will give a dilution of serum of 1/ 80 in Well A
through 1/ 10240 Well H.
5. Shake the plate gently to mix the contents thoroughly.
6. Incubate for 45-60 minutes at room temperature.( Keep the plate away from heat,
direct sunlight and any source of vibration).
7. Read results. Results are stable for 24hrs. if the plate is covered and the above
precautions are observed.
Table1: TPHA results
RESULTS
TEST CELLS
CONTROL CELLS
Strong Positive
Full cell pattern covering
the bottom of the well.
No agglutination tight
button
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Weak Positive
Cell pattern covers
approx. 1/3 of well
bottom
No agglutination tight
button
Indeterminate
Cell pattern shows a
distinctly open centre
No agglutination tight
button
Negative
Cells settled to a
compact bottom,
typically with a small
clear centre.
No agglutination tight
button
Positive reaction
Non-specific *
Non-specific absorption
Positive reaction
1. Add 100µl of test serum to a small tube then add 400µl of Control Cells.
Mix well and stand for 1 hour.
2. Centrifuge for 15 minutes at 1000 rpm and test the supernatant by the
qualitative method.
Note: The sample is now at 1/5, this should be taken into account when preparing the
dilutions. If the result is repeatedly non-specific the sample should be tested by another
method eg . FTA-ABS.
INTERPRETATION OF RESULTS
Strong positive reactions may show some folding at the edge of the cell mat.
When the Test well is positive , the Control well should be observed.
The Control cells should settle to a compact button. They should not be used as a
comparison for Non-Reactive serum patterns since the Control Cells will give a more
compact pattern than the Test Cells.
Agglutination in the Control well indicates the presence of non-specific agglutinins in the
sample, the test should be reported as INVALID. A serum that gives this result may be
absorbed using the Control Cells as detailed under Non-specific absorption. A doubtful
reaction with Test Cells should be reported as INDETERMINATE.
This result may indicate a low level of antibody in early primary syphilis or yaws.
This sample should be first retested in the qualitative test then a further sample should be
tested at a later date to determine whether or not there is a rising titre. It is also advisable
to perform another confirmation test (FTA-ABS) to complete the profile of the test
serum.
5.3 Widal Test ((agglutination test))
Introduction
119 | P a g e
The widal agglutination test for the diagnosis of the enteric fever, The patients' serum is
tested directly for antibody by entire a slide of tube method, These test are either qualitative
and semi quantitative.
The rapid slide test is used primarily as a screening procedure especially useful when large
numbers of sera must be examined. The tube test should be used to confirm positive result
obtained by the slide test.
The degree to which agglutination occurs depends upon the concentration of the antigen,
the amount of antibody present, the composition of the salt solution and the temperature.
Sera from normal patients may show positive agglutination with widal antigen due the
previously immunization, past infection, or the present of antibodies to related antigen. titer
detected as a result of active infection or recent immunization.
It is therefore, necessary to evaluate two or more serum samples, taken at 3 to 5 days
intervals after the onast of disease. a progressive increase in titers is the prime evidence of
recent infection or immunization.
Martials
1. antigens: Antigen are nonviable bacteria cells(salmonella)in 0.5% phenolized
saline and stained
2. Serological pipettes, 0.2 ml with 0.1 ml gradation. 0.1 ml capacity
3. Class agglutination slide with bow
4. Wooden applicator sticks
5. Physiological saline (0.85% NaCl)
6. Rotatory shaker (if available)
Procedure:
- Specimen collection: Collect 5-10 ml of blood in the sterile plain tube without
anticoagulant. Allow the blood to clot for 20-30 min, serum maybe kept for up
to 24 hours at 2.8 c. If sample to be kept for longer periods , it should be frozen,
do not refreeze one it thawed serum
The antigen should be allowed to warm to room temperature prior to use in test.
A- Slide method
1- deliver 0.08,0.04,0.02,0.01 and 0.005 ml of test serum on to separate ring, using 0.2
ml pipette
2- Shake the antigen vial in order to thoroughly mix the contents.
3- Add one drop of antigen to each circle, mix antigen and serum, using a new tooth
pick or applicator for each circle.
4- Rock the slide for 1-3 min (lock the kit procedure) or using rotatory shaker if
available.
120 | P a g e
5- The degree of agglutination observe in each ring is estimated as follow.
6- Dilution giving a 2+ agglutination is the end point or titer of that serum.
Note: storage kit: Widal kit, store at 2-8°C, antigen are ready for use as supplied
Measure titer:
The titer is the reciprocal of the highest final dilution procedure a 2 +agglutination
Table 2:Record agglutination
Degree of agglutination
100%
back ground clear to slightly hazy
4+
75%
back ground slightly cloudy
50%
back ground moderately cloudy
2+
25%
back ground cloudy
1+
3+
None
negative
Table 3: Determination of antibody titer
The titer is reciprocal of the highest dilution producing (2+) agglutination
Serum ml
correlation dilution
0.08
1:20
0.04
1:40
0.02
1:80
0.01
0.005
1:160
1:320
121 | P a g e
Table 4: Example of slide test are shown in the following table
Slide test results
Slide test result
Serum ml
readying
0.08
0.04
0.02
0.01
0.005
1:20
1:40
1:80
1:160
1:320
dilution
Serum
titers or
endpoint
Agglutination reaction
Serum No
1
++++
++++
++++
++
-
1:160
Serum No
2
++++
+++
++
-
-
1:80
Serum No
3
+++
++
++
_
_
1:80
Note:
1-If auto-agglutination of antigen is suspected (Show by low power microscope) the
vial antigen is unstable and should be discarded.
2-patints, occasionally fail to develop any serum agglutinin
The chart below give approximate indication of the significant of serum titers.
Table 5: indication of significant serum titers
Serum agglutinins
Disease
Febric antigen
widal antigen
appear
maximum
titer and
significance
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Typhoid fever
Para typhoid
fever
salmonella
infection
Salmonella O group
D
Salmonella O group
A
Salmonella O group
B
7-10
Days
3-5 week
1:80 (in early
stage)=suspicious
1:160 and rising= strong
suspicious
Those characterized by
prolonged fever and
typhoid like symptoms
present antibodies of
titers similar to above
Note:
1-The minimal titers defined as positive of the O typhoid antigen and H antigen must be
determined for individual geographic areas.
2-Cross-reaction occur with both non Styphi group dsalmonella and salmonella from
other groups
3-A single elevated titer for (O) equal to or greater than (1:320) or H equal to or greater
then (1:640) is considered positive.
5.4 Rose Bengal Test for Brucellosis
Introduction
Human brucellosis –undulant or Malta fever:
Brucellosis is systemic infection in which any organ or system of body can be involved.
Human become infected by three primary routes
1-ingestion of infected unpasteurized animal milk products
2-inhaltion of infected aerosolized particles
3-Direct contact with infected animal parts through rupture of skin and mucous
membrane including the conjunctiva.
Brucellosis is one of the most commonly reported bacterial infection acquired in
laboratory , within laboratory aerosolizing is primary mechanism of the transmission, so
it is required biosafety level 3 precaution for isolation in Lab .
Most species of pathogenic for human are:
1- Br abroteus from cows
2- Br melitens from goats and sheep
3- Br suis from pigs (USA)
4- Br canis from dog
Definitive diagnosis of brucellosis require isolation of the organism in culture of blood ,
bone marrow or other tissues.
123 | P a g e
the cultures , should be incubated in 5% to 10 % CO2 in humidified atmosphere incubated
for 3 weeks before discarded as negative.
The mean way of diagnosis of brucellosis is by serological test.
Become of the difficulty of isolating the organism (serological test) the serum agglutination
test (SAT) is widely used and detected antibodies for Br abroteus and Br melitensis.
Rose Bengal test (agglutination test) for brucellosis it is remain an inexpensive sensitive
and specified test for acute case , but it is not useful in following up of patients or in
diagnosis relapses .
Principle
The rose Bengal an agglutination test reaction used for the diagnosis of brucellosis.
The patients’ serum is tested directly for antibody by either a slide or tube method.
The test are qualitative and semi qualitative.
A number of antigens are available commercially for use is the rapid slide test.
The test is performed by mixing serum and antigen in ruled squired on a glass slide.
This test is widely used and detected antibodies to Brucellosis aborteas, Br. Melitensis.
In case of a cute infection a titer of 1:160 or greater in rapid slide method are consider
diagnosis if this result fits the clinical and epidemiologic findings.
Materials:
1. Antigen Brucellosis aborteas, nonviable. Bacterial cells in 0.05% phenolized
saline and stained.
2. Serological pipettes 0.2 ml with 0.1 ml graduation. 0.1 ml capacity
3. Class agglutination slide with row
4. Wooden applicator sticks
5. Physiological saline(0.85% NaCl)
6. Rotator shaker if available
Procedure
Specimen collection
Collect 5-10 ml of blood in the sterile plain tube without anticoagulant. Allow blood
to clot for 20-30 mint, serum may be kept for up to 24 hours at 2-8 C° e . If samples
to be kept for longer period, it should be frozen don’t refreeze once themed serum
A1.
2.
3.
The antigen should be allow to warm to room temperature prior to use in test
Slide method.
delivers 0.08,0.04,0.02,0.01 and 0.005 ml of test serum on to separate ring using 0.2
ml pipette.
Shake the antigen vial in order to thoroughly mix the contents.
Add one drop of antigen to each circle mixer antigen and serum , using a new tool
pick or applicator for each circle.
124 | P a g e
4. Rock the slide for 1-3 min (lock the kit procedure) or using rotatory shaker if
available.
5. The degree of agglutination observe in each ring is estimated as follow.
Record agglutination
Degree of agglutination
100%
back ground clear to slightly hazy
75%
back ground slightly cloudy
50%
back ground moderately cloudy
25%
back ground cloudy
None
4+
3+
2+
1+
negative
Note: Storage kits Rose Bengal kit store at 2-8C° antigen are ready for use as supplied
Table 6: Determination of antibody titer
The titer is reciprocal of the highest dilution producing (2+) agglutination
Serum Ml
correlation dilution
0.08
1:20
0.04
1:40
0.02
1:80
0.01
0.005
1:160
1:320
The serum dilation giving a 2+ agglutination is the end point or titer of the serum.
Measure titer is reciprocal of the highest final dilution producing (2+) agglutination
Example of rose bagel test result are shown in the flowing table
Table 7: Examples of Rose Bengal results
Slide test result
0.08
0.04
0.02
0.01
0.005
Serum
titers or
endpoint
125 | P a g e
Serum
(ml)
readying
1:20
1:40
1:80
1:160
1:320
dilution
Agglutination reaction
Serum No
1
++++
++++
++++
++
-
1:160
Serum No
2
++++
+++
++
-
-
1:80
Serum No
3
+++
++
++
_
_
1:80
Limitation of the procedure
1- The serum agglutination test SAT measure both antibodies IgG and IgM in serum of
patient
2- After cure the antibodies for brucellosis, usually persistent for more than one year
3- Use 2-mercaptoethanol (detect IgG only) to identify chronic or relapse Brucellosis
cases.
4- Any titer high than 1:40 with 2-mercaptoethanol lgG indicative active infection (i.e.
relapses or chronic infection).
5- A single titer of greater than 160 or more is considered significant.
6- Prozone phenomena may give false negative result which is possible because of the
high antibody concentration so serum dilution is recommended to avoid this.
7- False positive reaction may result from cross-reaction with antibodies of cholera,
yersinia or francisella.
8- False negative and false positive reaction can be avoided by routinely dilution the
serum beyond at 1:320
126 | P a g e
Module 3
Session 2: CRP Test, ASO latex slide agglutination Test, RF, Toxoplasmosis Test,
Infectious mononucleosis Test, Rota Virus Latex Test, and Pregnant Test
Objectives
At the end of this session participants will be able to:
1. Understand CRP Test
2. Understand ASO latex slide agglutination Test
3. Understand RF
4. Understand Toxoplasmosis Test
5. Understand Infectious mononucleosis Test
6. Understand Rota Virus Latex Test
7. Understand Pregnant Test
Trainers Preparation:




Review the reading material and the session plan.
Prepare the presentation as appropriate and as recommended in the method
column of the session plan, or write the information on a flipchart or board where
all participants can see it.
Prepare copies of the reference materials/handouts and exercises.
Arrange the training room.
Methods and activities

Mini-lecture, question & answers, discussion-lecture
Evaluation/assessment
Questions and answers, trainer’s observation
Estimated Time
190 minutes
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Session plan -2-
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Objectives
3.2.1. Understand CRP
Test
3.2.2.Understand ASO
latex slide agglutination
Test
3.2.3. Understand RF
Content





Principle
Material
Reagent preparation
Procedure
Results and
interpretation


Principle
Material and
reagents
Procedure









3.2.4. Understand
Toxoplasmosis Test




Principle
Material
Specimen collection
and preparation
Reagent preparation
Assay procedure
Results and
interpretation
Qualitative method
Semi-Quantitative
method
Principle of test
Materials
Method
Manual wash
procedure
Methodology
Mini-lecture
25 minutes
Question & answers
20 minutes
Mini-lecture
30 minutes
Discussion- lecture
45 minutes
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
3.2.5. Understand
Infectious mononucleosis
Test
3.2.6. Understand Rota
Virus Latex Test
Reagents of
infectious
mononucleosis test
Methods
Results and
interpretation in
reports
Question & answers




Principle
Materials
Method
Result and
interpretation
Question & answers

Rapid test for
detection of human
chorionic
gonodotrophin in
urine
Direct hGG latex
pregnancy kit


3.2.7.Understand Pregnant
Test

20 minutes
20 minutes
Discussion- lecture
30 minutes
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5.5 C-REACTIVE PROTEIN ( CRP) TEST
PRINCIPLE
C-Reactive Protein (CRP) in patient’s sera has been found in association with acute
infections, necrotic conditions and a variety of inflammatory disorders. There is a strong
correlation between serum levels of CRP and the onset of the inflammatory process.
Monitoring the levels of CRP in patient’s sera indicates the effectiveness of treatment and
the assessment of patient recovery.
CRP latex particles are coated with antibodies to human CRP. When the latex suspension
is mixed with serum containing elevated CRP levels on a slide, clear agglutination is seen
within 2 minutes.
MATERIAL
1- Suspension of polystyrene latex ( approximately 1%) particles coated with AntiCRP antibodies.
2- Positive Control. Serum containing CRP antibodies.
3- Negative Control. Serum free of CRP antibodies.
4- Stirrers
5- Plastic slide
6- Micro-pipettes capable of dispensing 50μl.
7- Isotonic saline (0.9% NaCI)
REAGENT PREPARATION
o
o
All reagents should be brought to room temperature (20 C to 25 C) and mixed gently to
resuspend latex prior to use. Do not induce foaming.
The test slide should be thoroughly cleaned before use as traces of detergent or prior
specimen may affect the result.
Recommended cleaning procedure:
1. Used cards must be immediately immersed in a disinfectant solution. Follow
disinfectant manufactures guidelines.
2. The reaction circles must be physically rubbed with non-abrasive material to ensure
removal of possible adhering particles.
3. Thoroughly rinse in purified water.
4. Allow reaction card to dry.
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5. Spray cards with a 70% alcohol solution.
6. Allow the alcohol to evaporate prior to re-use.
PROCEDURE
Qualitative Method
1. Allow kit reagents and patient serum to come to room temperature.
2. Transfer one drop (50μl) of patient’s serum to the test circle on the slide.
3. Shake the latex reagent, then using the dropper provided, add one drop of suspension
to the test circle.
4. Mix the drops using a disposable stirrer ensuring coverage of the test circle with the
mixture.
5. Gently and evenly, rock and rotate the test slide for 2 minutes whilst examining the test
slide for agglutination.
Semi Quantitative Method
1. Using isotonic saline prepare serial dilutions of the patients serum (1/2, 1/4, 1/8, 1/16,
1/32, 1/64 and so on)
2. Transfer one drop (50μl) of each serum dilution to the test circle on the slide.
3. Shake the latex reagent, then using the dropper provided, add one drop of suspension
to the test circle.
4. Mix the drops using a disposable stirrer ensuring coverage of the test circle with the
mixture.
5. Gently and evenly, rock and rotate the test slide for 2 minutes whilst examining the test
slide for agglutination.
RESULTS AND INTERPRETATION
Examine the test slide under a strong light source after 2 minutes. Kit controls or known
level value samples should be tested with each test run. The kit negative control should
give a negative result after 2 minutes. The kit positive control should give a positive
result at a titre of 1/4 +/- one double dilution after 2 minutes. If levels of controls or users
known samples do not give expected results, test results must be considered invalid.
Qualitative Method
A positive result is indicated by the obvious agglutination pattern of the latex, in a clear
solution. A negative result is indicated by no change in the latex suspension on the test
slide.
Latex CRP kit has a detection limit of 6mg/litre of CRP in the patient’s serum. Positive
results will be obtained at a CRP serum concentration above 6mg/litre and negative
results will be obtained at 6mg/Litre and below.
Semi-Quantitative Method
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The serum CRP concentration can then be calculated approximately by multiplying the
dilution factor (i.e 2, 4, 8 or 16) by the detection limit, i.e. 6, to give the number of
mg/Litre concentration e.g. if the agglutination titre appears at 1/8 the approximate serum
CRP concentration is 8 x 6 = 48 mg/Litre.
Titres of 500 mg/Litre have been detected with CRP with no prozone ( hook ) effect.
5.6 ASO latex slide agglutination test
Principle
Streptococcal infections are very common in all populations, and a high percentage of
people will have antibodies against streptococci. The b-haemolytic group A streptococci
produce two haemolysins: oxygen-labile streptolysin O and oxygen-stable haemolysin S.
Only reduced (non-oxidized) streptolysin O is immunogenic and is used for the test. The
antistreptolysin O test is based on the fact that patients with Streptococcus pyogenes
(group A streptococcal) infections develop antibodies that inhibit the haemolytic activity
of streptolysin O. The antibodies are usually long-lasting and a single increased titre
is not an indication of a current infection. Only a fourfold or greater rise in titre on
successive serum samples taken 10–14 days apart should be considered indicative of
recent infection. This test is mainly used in the diagnosis of acute rheumatic fever, acute
glomerulonephritis and other post-streptococcal
diseases.
There are two types of commercial antistreptolysin O test kits:
The ASO latex slide agglutination test is used to screen sera to identify those with raised
ASO titres (200 IU or higher).
The ASO tube test is a haemolysis inhibition test that is used to determine ASO antibody
titre in serum samples that are positive in the ASO latex slide agglutination test. Atitre of
less than 50 IU does not confirm the diagnosis of acute rheumatic fever.
Materials and reagents
1- Disposable cards, with 6 wells each Disposable dropper
2- Positive control serum
3- Sensitized latex reagent (with streptolysin O)
4- Applicator sticks
Procedure
1- Dilute the serum 1: 20. (Or according to kit literature).
2- Place 1 drop of the serum solution in a well on the disposable card.
3- Use a new dropper to add 1 drop of sensitized latex reagent.
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4- Use an applicator stick to mix the two drops and spread them over the entire well.
5- Examine for agglutination within 2 minutes.
 A positive reaction appears as a fine agglutination within 2 minutes.
 A negative reaction shows no agglutination.
5.6 Rheumatoid Factor (RF)
PRINCIPLE
Rheumatoid Factor (RF) is found in sera of patients with Rheumatoid Arthritis and is
believed to be IgM antibodies directed against the patient’s own immunoglobulin G.
RF is a rapid latex agglutination test kit for the detection of (RF) in human serum. RF
latex particles are coated with specially purified human gamma globulin, When the latex
suspension is mixed with serum containing elevated RF levels on a slide, clear
agglutination is seen within 2 minutes. RF has a detection limit of 8 IU/ml of RF in the
patient serum .
Material
1- Suspension of Polystyrene latex particles ( approximatly 1.25% ) coated with
suitably modified Fc fraction of IgG in stabilizing buffer.
2- Positive Control. Serum containing Rheumatoid Factor antibodies. Working
Strength.
3- Negative Control. Serum free of Rheumatoid Factor antibodies.
4- Stirrers
5- Plastic slide
6- Micropipettes capable of dispensing 50 μl.
7- Isotonic saline (0.9% NaCI)
SPECIMEN COLLECTION AND PREPARATION
Obtain a sample of venous blood from the patient and allow a clot to form and retract.
Centrifuge clotted blood sample and collect clear serum. Fresh serum samples are
required.
Do not use haemolysed, contaminated or lipaemic serum for testing as this will adversly
affect the results.
o
o
Serum may be stored at 2 C to 8 C for up to 48 hours prior to testing. If longer storage is
o
required, store at –20 C for up to 6 weeks. Thawed samples must be mixed prior to
testing.
Do not repeatedly freeze-thaw the specimens as this will cause false results.
REAGENT PREPARATION
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o
o
All reagents should be brought to room temperature (20 C to 25 C) and mixed gently
prior to use. Do not induce foaming.
The test slide should be thoroughly cleaned before use as traces of detergent or prior
specimen may affect the result.
Recommended Cleaning procedure:
1- Used cards must be immediately immersed in a disinfectant solution. Follow
disinfectant manufactures guidelines.
2- The reaction circles must be physically rubbed with non-abrasive material to
ensure removal of possible adhering particles.
3- Thoroughly rinse in purified water.
4- Allow reaction card to dry.
5- Spray cards with a 70% alcohol solution.
6- Allow the alcohol to evaporate prior to re-use.
ASSAY PROCEDURE
Qualitative Method
1. Allow kit reagents and patient serum to come to room temperature.
2. Transfer 50μl of patient’s serum to the test circle on the slide.
3. Shake the latex reagent, then using the dropper provided, add one drop of suspension
to the test circle.
4. Mix the drops using a disposable stirrer ensuring coverage of the test circle with the
mixture.
5. Gently and evenly, rock and rotate the test slide for 2 minutes whilst examining the test
slide for agglutination.
Semi Quantitative Method
1. Using isotonic saline prepare serial dilutions of the patients serum (1/2, 1/4, 1/8, 1/16,
1/32, 1/64 and so on)
2. Transfer 50μl of each serum dilution to the test circle on the slide.
3. Shake the latex reagent, then using the dropper provided, add one drop of suspension
to the test circle.
4. Mix the drops using a disposable stirrer ensuring coverage of the test circle with the
mixture.
5. Gently and evenly, rock and rotate the test slide for 2 minutes whilst examining the test
slide for agglutination.
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RESULTS AND INTERPRETATION
Examine the test slide under a strong light source after 2 minutes. Kit controls or known
level value samples should be tested with each test run. The kit negative control should
give a negative result after 2 minutes. The kit positive control should give a positive
result at a titre of 1/4 +/- one double dilution after 2 minutes. If levels of controls or users
known samples do not give expected results, test results must be considered invalid.
QUALITATIVE METHOD
A positive result is indicated by the obvious agglutination pattern of the latex, in a clear
solution. A negative result is indicated by no change in the latex suspension on the test
slide.
Positive results will be obtained at a RF serum concentration of 8 IU/ml or more and
negative results will be obtained at a RF concentration below 8 IU/ml.
SEMI – QUANTITATIVE METHOD
The serum RF concentration can then be calculated approximately by multiplying the
dilution factor (i.e 2, 4, 8 or 16) by the detection limit, i.e. 8, to give the number of IU/ml
concentration e.g. if the agglutination titre appears at 1/8 the approximate serum RF
concentration is 8 x 8 = 64 IU/ml.
Titres of 1074 IU/ml have been detected with Avitex RF with no prozone ( Hook ) effect.
5.7 Toxoplasmosis Test
Toxoplasma IgG ELISA
Introduction
Toxoplasma gondii is an intracellular protozoan parasite with a worldwide distribution.
Although cats are the definitive host, human infection results from ingestion of
contaminated soil, careless handling of cat litter, ingestion of raw or undercooked meat or
transmission from mother to foetus through the placenta. When a seronegative woman
becomes infected with T. gondii during pregnancy, the organism is often transmitted to
the foetus. Infection during the first trimester may lead to spontaneous abortion, stillbirth,
or overt disease in the neonate.
The Toxoplasma IgG kit is a rapid ELISA designed for the qualitative or quantitative
detection of IgG antibodies to Toxoplasma gondii in human serum.
Principle of the test
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Diluted serum or plasma specimens (1:100) are incubated for 20
minutes to allow specific antibodies to T. gondii to bind to the antigencoated wells. After
washing away unbound antibodies and other serum constituents, T. gondii specific IgG is
detected using rabbit anti-human IgG conjugated to horseradish peroxidase. After 20
minutes incubation, unbound conjugate is removed by washing, and TMB enzyme
substrate is added for 10 minutes. A blue colour develops if antibodies to T. gondii are
present. Addition of stop solution gives a yellow colour and the optical densities of
controls, standard(s) and samples are measured using a microplate reader.
Materials
1. Microplate 96 wells in 12 X 8 break-apart strips, pre-coated with T.gondii purified
antigens enriched for P30 antigen
2. Reagent 1: Sample Diluent 100mM Tris-buffered saline, pH 7.2 with antimicrobial
agent, 10ml, (blue), concentrate (x10)
3. Reagent 2: Wash Buffer 100mM Tris-buffered saline with detergent, pH 7.2, 100 ml,
concentrate (x10)
4. Reagent 3: Conjugate (peroxidase conjugated rabbit anti-human IgG), 12 ml, (red),
ready to use
5. Reagent 4: TMB Substrate, 12 ml, ready to use
6. Reagent 5: Stop solution, 12 ml, ready to use
7. Standards1: (for quantitative assays), 15 IU/ml, 50 IU/ml; 150 IU/ml (blue), ready to
use
8. Standard1: (for qualitative assays) 8 IU/ml, (yellow), ready to use
9. Positive control1: 100 IU/ml (red), 1ml, ready to use
10. Negative control: 1 ml (green), 1ml, ready to use
11.Distilled or de-ionised water
Equipment
12345-
10mm X 60mm tubes for dilution,
pipettes 10μl, 100μl, 1000μl;
repeating dispenser 100μl,
microplate reader with 450nm filter,
microplate washing device.
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Samples
Only freshly drawn and properly refrigerated sera or plasma should be used in this assay.
Avoid haemolysed, lipemic or bacterial contaminated sera. Sera should be stored at 2-8ºC
for no longer than 5 days. If delay in testing is anticipated, store test sera at –20ºC. Avoid
multiple freeze-thaw cycles.
Method
Ensure that all materials are at room temperature before beginning the procedure. We
recommend that the standards and the controls are always run in duplicate. Samples may
be run singly or in duplicate.
1. Assemble the number of strips required for the assay.
2. The sample diluent X10 concentrate contains 0.09% sodium azide as preservative.
Prepare sufficient working strength diluent for the assay run. However, if the working
strength diluent is to be stored for more than 1 week, add sodium azide (0.9g/L). Store
unused sample diluent concentrate and dilute sample diluent at 2 – 8oC. Dilute the
Sample Diluent (Reagent 1) 1:9 in distilled water to make sufficient buffer for the assay
run e.g. add 10ml sample diluent concentrate to 90 ml water.
3. Dilute patient samples 1:100 (e.g. 5μl serum plus 0.5 ml diluent). It is important to
dispense all samples and controls into the wells without delay. Therefore ensure that all
samples are ready to dispense.
4. For qualitative determinations, dispense 100 μl of the negative control, the 8 IU/ml
standard, the positive control and the diluted patient sample into the wells. For
quantitative determinations, use sample diluent as 0 IU/ml and additionally dispense the
15 IU/ml, 50 IU/ml and 150 IU/ml standards.
5. Place the strips into the incubation bag provided and incubate for 20 minutes at room
temperature. During all incubations, avoid direct sunlight and close proximity to any heat
sources.
6. Dilute the Wash Buffer (Reagent 2) 1: 9 in distilled water to make sufficient buffer for
the assay run e.g. add 50ml wash buffer concentrate to 450ml water. The diluted wash
buffer is stable for two months at 2 - 8°C.
7. After 20 minutes, decant or aspirate the well contents and wash the wells 3 times using
an automatic plate washer or the manual wash procedure (see below). Careful washing is
the key to good results. Blot the wells on absorbent paper before proceeding. Do not
allow the wells to dry
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Out .
Manual Wash Procedure:
Empty the wells by inversion. Using a multi-channel pipette or wash bottle, fill the wells
with ash buffer. Empty by inversion and blot the wells on absorbent paper. Repeat this
wash process two more times.
8. Dispense 100μl of Conjugate (Reagent 3) into each well.
This reagent is colour coded red. Keep all pipettes and other equipment used for
Conjugate ompletely separate from the TMB Substrate reagent! Incubate the wells for 20
minutes in the incubation bag at room temperature.
9. After 20 minutes, discard the well contents and carefully wash the wells four times
with wash buffer. Ensure that the wells are completely washed. Blot the microplate on
absorbent paper to remove final drops of wash fluid. Do not allow the wells to dry out.
10. Using a repeating dispenser, rapidly dispense 100μl of TMB Substrate (Reagent 4)
into each well. Incubate the plate for 10 minutes.
11. Add 100μl of Stop Solution (Reagent 5) to each well. To allow equal reaction times,
the stop solution should be added to the wells in the same order as the TMB Substrate.
12. Read the optical density in a microplate reader within 10 minutes.
Interpretation
Qualitative determinations
Negative samples: OD < 8 IU/ml OD
Positive samples: OD >/= 8 IU/ml OD
Quantitative determinations
Plot the optical densities of the standards against their respective concentrations. Draw a
line to join the points. Read the concentrations of unknowns from this graph.
Concentrations below 8 IU/ml are considered negative; concentrations above 8 IU/ml are
considered positive for anti-toxoplasma IgG. A negative result indicates no current or
previous infection with T.gondii. Such individuals are presumed to be susceptible to
primary infection. However see Limitations below. A positive result indicates a current or
previous infection with T. gondii.
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5.8 Infectious Mononucleosis test
Principle
Is a qualitative determination of the nonspecific heterophile antibodies that result from
primary infection by Epstein – Barr Virus and causing the clinical presentation of
Infectious mononucleosis.
The IM- Latex is a slide agglutination test also can get semi-quantitative detection of
Heterophile antibodies . Latex particles coated with antigenic extract of beef erythrocyte
membranes and are agglutinated when mixed with samples containing IM heterophile
antibodies.
Materials and reagents Preparedness:
 Reagents
Table 8: Reagents of Infectious Mononucleosis Test
Latex
Latex particles coated with antigenic extract of beef
erythrocytes membranes , phosphate buffer, pH 7.2.
Sodium azide , 0,95 g/L.
Control +ve
Human serum with anti – IM antibodies titer ≥ 1/4 .
Sodium azide , 0.95g/L
Control -ve
Animal serum . Sodium azide , 0.95g/L
Disposable slides
Storage: when stored tightly closed at 2-8 oC do not freeze. (according to kit
literature)

Samples : Fresh serum . Stable 7 days at 2-8 oC or 3 months at – 20 oC. Samples
with presence of fibrin should be centrifuged. Do not use highly hemolized or
lipemic samples.
 Equipment :
1- Mechanical rotator with adjustable speed at 80-100 r.p.m.
2- Centrifuge.
3- Automatic micropipette
Methods:
 Qualitative Method :
1- Allow reagents and samples to reach room temperature.
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2- Place 50 ul of the sample and one drop of each Positive and Negative controls into
separate circles on the slide test.
3- Swirl the IM – Latex reagent gently before using and add one drop ( 50ul) next to
the samples to be tested.
4- Mix the drops with stirrer , spreading them over the entire surface of the circle .
Use different stirrers for each sample.
5- Place the slide on a mechanical rotator at 80-100 r.p.m. for 2 minutes .False
positive results could appear if the test is read later than two minutes.
 Semi quantitative method:
1- Make serial two ford dilutions of the sample in 9 g/L saline solution.
2- Proceed for each dilution as in the qualitative method.
Results and Interpretations in reports:
Examine macroscopically the presence or absence of visible agglutination immediately
after removing the slide from the rotator. The presence of agglutination indicates a titer ≥
1/28 of anti IM heterophil antibodies by the Davidson method.
The titer in the semi- quantitative method is defined as the highest dilution showing a
positive result.
The following are the limitations of the test:
1- False positive results may be obtained in some geographical areas where the horse
serum is used as a prophylactic measure (vaccination). Also this could be seen in patients
with leukemia, Burkitts Lymphoma, Pancreatic carcinoma, viral hepatitis, CMV
infections and others.
2- False negative results also encountered and in such Patients either repeat testing after
interval of several days to get a high heterophil antibody titer or look for specific
antibodies against viral structural antigens like anti VCA IgM .
Quality Control:
Positive and Negative controls are recommended to monitor the performance of the
procedure, as well as the comparative pattern for a better result interpretation.
5.9 Rota Virus Latex Test
Principle of the method and clinical significance:
Rapid latex agglutination assay for the detection of rotavirus in faecal samples this virus
has been shown to be principle causative agent of gastro-enteritis. Patients may harbor up
to 108 virus particles per gram of faeces.
Materials and reagents Preparedness:
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
Material insert in kit: Test reagent latex, control negative ,Positive bovine rotavirus control inactivated ( ready to use ),5 x concentrated extraction buffer pH
7.2,Agglutination slides ,pipette
 Specimen collection and storage: Prepare a 10% suspension of the faecal sample
by adding 0.1 g /0.1 ml of sample to 1 ml of extraction buffer in screw capped
vial. Mix well and stand in room temperature for 2 minutes. Specimen may be
stored 24 hours at the time of the test .If storage is more than 24 hour the
specimen should be stored at -20 oC in freezer .
 Safety warnings: Do not pipette by mouth, use disinfectant 70% alcohol, waste
must be treated as biohazardous and incinerated.
Method:
123456-
Add 10 ml of extraction buffer to 40ml de-ionize water.
Add 2 gms of faecal sample to 2 ml of extraction buffer.
Mix well on vortex mixer.
Stand at room temperature for 10 minutes.
Centrifuge test sample for 10 minutes about 800 g.
Place one drop of the supernatant from the faecal sample onto the circle of test
slide and use the 2 circle for control reagent.
7- Add one drop of control latex reagent next to the drop of faecal sample on circle
1.
8- Add one drop of test latex reagent next to drop of faecal sample on circle 2.
9- Spread the control latex reagent and test latex reagent in circle 1, 2 by disposable
pipette.
10- Gently tilt the test slid backward and forward for 2 minutes.
Results and Interpretations in reports:
1- Positive result is indicated by the visible agglutination of the latex particles which
occur within few seconds.
2- Negative result is indicated by milky appearance without any visible aggregation
of the latex particles.
5.10 PREGNANCY TEST
5.10.1 Rapid test for detection of human Chorionic Gonadotrophin in Urine
Human chorionic gonadotrophin (hCG), a glycoprotein hormone secreted by viable
placental tissue during pregnancy, is excreted in urine approximately 20 days after the
last menstrual period. The levels of hCG rise rapidly reaching peak levels after 60-80
days. The appearance of hCG in urine soon after conception and its rapid rise in
concentration makes it an ideal marker for the early detection and confirmation of
pregnancy. Test kit utilizes the principle of Immunochromatography, a unique two-site
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immunoassay on a membrane. As the test sample flows through the membrane assembly
within the test device in place of the dipstick, the coloured monoclonal anti-hCGcolloidal gold conjugate complexes with the hCG in the sample. This complex moves
further on the membrane to the test region where it is immobilised by the monoclonal
anti-hCG coated on the membrane, leading to the formation of a pink coloured line which
confirms a positive test result. Absence of this coloured line in the test region indicates a
negative test result. The unreacted conjugate, unbound complex, if any, and the colloidal
gold conjugated rabbit IgG moves further along the membrane and are subsequently
immobilised by the goat anti-rabbit IgG coated on the control region forming a pink line.
This control line serves to validate the test results.
SPECIMEN COLLECTION AND PREPARATION
Urine samples collected at any time may be used, however, it is recommended that to
maximize hCG concentration, the first voided morning specimen should be used. The
urine should be collected in a clean dry container (plastic or glass) which must be free
from detergent. Urine specimens should be as fresh as possible and it is preferable to test
within 24 hours of collection. The sample may be stored for longer periods (72 hours)
prior to use, if stored, store at 2oC to 8oC.
Filtration or centrifugation is generally not necessary for urine used in this test, however,
if a sample is very turbid, centrifugation or filtration may be necessary. (The use of
supernatant from turbid samples allowed to sediment naturally prior to use may negate
the need for sample preparation – this does not affect the hCG concentration.)
REAGENT PREPARATION
Devices and samples should be brought to room temperature (20 to 25) °C and mixed
gently prior to use.
In case the pouch has been stored at 4°C to 8°C, allow at least 30 minutes for the device
to come to room temperature. Check the color of the desiccant. It should be blue. If it has
turned colorless or faint blue, discard the device and use another device.
ASSAY PROCEDURE
1. Open the pouch and remove the device. Once opened, the device must be used
immediately.
2. Dispense two drops of urine specimen into the sample well „S‟ using the dropper
provided.
3. Read the results at the end of fifteen minutes for serum or five minutes for urine.
RESULTS AND INTERPRETATION
Negative: Only one colored line appears on the control region „C‟ only
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Positive: A distinct colored line appears on the control region „C‟ and on the test region
“T”.
Figure 14: Results of PT
The test should be considered invalid if no line appears. Repeat the test with a new
device.
Depending on the concentration of hCG in the specimen, positive results may appear as
early as 30 seconds. Negative results must be confirmed only at the end of fifteen minutes
for serum or five minutes for urine.
5.10.2 DIRECT hCG LATEX PREGNANCY KIT
Detectable levels of Chorionic Gonadotropin (hCG) in urine start at 5 mIU/ml during the
first week of gestation and rises to 100,000 mIU/ml at 2 to 3 months. The hCG level
doubles approximately every 2.2 days during the first trimester1. Values decline from
10% to 15% of peak concentrations during 2nd and 3rd trimesters2.
Direct hCG Latex kit contains one reagent of latex particles coated with monoclonal
antibodies to hCG. The reagent is mixed with the urine samples.
MATERIALS
1. Latex reagent.
2. Positive and negative controls.
3. Reaction slide and stirring sticks.
SPECIMEN COLLECTION AND PREPARATION
Generally, the first morning urine contains the highest concentration of the hCG hormone
and therefore, it is more recommended for testing. However, urine collected at other
periods can also be used. The urine in this case should have been kept in 2-8°C and used
within 72 hours from collection time.
PROCEDURES
1. Bring reagents to room temperature.
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2. Place 50μl of pa�ent urine, one drop positive and one drop negative controls into
different circles of the slide.
3. Add one drop of latex reagent directly to each sample.
4. Mix using the supplied sticks and spread the mixture over the entire circle.
5. Gently rock the slide. Agglutination may be observed after two minutes. Direct
light source may help to observe the results.
READING THE RESULT
Presence of agglutination within two minutes indicates positive reaction.
Lack of agglutination within two minutes indicates negative reaction.
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Module four: Bacteriology
Module Objectives:
At the end of this module the participant will be able to:
1. Apply bacteriological tests
Modules Sessions
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Session 1: General urine examination and general stool examination
Session 2: Urine culture, throat swab, and nasal swab
Session 3: Smart tests, preparation of media, and staining smears
Evaluation/ Assessment
Questions and answers, participants’ summaries, trainer’s evaluation
Estimated Training Time
10 hours
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Module 4
Session 1: General urine examination and general stool examination
Objectives
At the end of this session participants will be able to:
1. Explain macroscopical examination of urine
2. Detect and estimate glucose and protein in urine
3. Detect bilirubin in urine
4. Detect ketones bodies in urine
5. Detect abnormal elements in urine
6. Explain macroscopical examination of stool
7. Explain microscopical examination of stool
Trainers Preparation:
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

Review the reading material and the session plan.
Prepare the presentation as appropriate and as recommended in the method
column of the session plan, or write the information on a flipchart or board where
all participants can see it.
Prepare copies of the reference materials/handouts and exercises.
Arrange the training room.
Methods and activities

Mini-lecture, discussion, brain storming, question & answers, discussion-lecture
Evaluation/assessment
Questions and answers, trainer’s observation
Estimated Time
230 minutes
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Session plan -1-
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Objectives
4.1.1.Explain
macroscopical examination
of urine
4.1.2.Detect and estimate
glucose and protein in
urine
Content
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Introduction
Appearance
Odor
Reaction ( PH )
Principle
Method

Detection of
glucose in urine
Detection and
estimation of
protein in urine



4.1.3.Detect bilirubin in
urine

Introduction
Clinical
significance
Method
Methodology
Mini-lecture
30 minutes
Discussion
30 minutes
Brain storming
30 minutes
Question & answers
4.1.4.Detect ketones bodies
in urine
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
Introduction
Principles
Materials
Method
Result
30 minutes
Discussion-lecture

Microscopical
examination

Colour
4.1.5.Detect abnormal
elements in urine
4.1.6.Explain
60 minutes
Question & answers
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macroscopical examination
of stool
4.1.7.Explain
microscopical examination
of stool
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Consistency
Pus
Mucus
Method
Intestinal protozoa
Identification of
cysts
Importance of cysts
20 minutes
Mini-lecture
30 minutes
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6. Bacteriology
6.1 General urine examination
Introduction:
The examination of urine provides a wide variety of useful medical information
regarding the disease involving the kidney and lower urinary tract . both functional
(physiologic ) and structural (anatomic disorders of the kidney and lower urinary tract
my be elucidated . and diagnostic .monitoring, we believe laboratory urine tests will
remain an essential part of clinical medicine
The basic routine analysis consist of 4 parts : specimen evaluation , gross (physical)
examination , chemical screening , and sediment examination
Clinical significance
Examination of urine is a fundamental investigation in patients in whom kidney disorders
or infections of the urinary tract are suspected. There are also many patients who exhibit
no clinical symptoms, but in whom previously unrecognized urinary tract infections can
be diagnosed by urine examination
Collection of urine specimens
Containers for the collection of urine should be wide-mouthed, clean and dry. And we
can also be used the adhesive bag for infants .If the urine specimen has to be transported
for any length of time it should contain an appropriate preservative to prevent bacterial
overgrowth or hatching of viable ova The specimen can be preserved for microscopic
examination of the deposit by adding 8–10 drops of formaldehyde, 10% solution per 300
ml of urine. Urine preserved in this way is not suitable for other tests.
Urine to be examined under the microscope must be freshly passed into a clean dry
vessel. A midstream urine specimen is the most useful. Urine stored in a refrigerator may
contain an excess of precipitated salts and will not be suitable for microscopy
principle
the elements of routine urinalysis include macroscopical examination the evolution of
physical characteristics ( color, odor, and specific gravity ) pH and chemical
examination ( protein , glucose. Keton bodies, bilirubin, urobilinogen. Bile pigment and
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bile salt) and microscopic inspection of centrifuge sediment for R.B.C pus, cast. crystal.
etc..
Material( glassware & Equipment):
1- centrifuge
2- Microscope
3- Urine cup
4- conical centrifuge tube
5- slides
6- cover slide
7- Watch glasses
8- Dropper
9- Forceps
10- Indicator paper of limited pH range: for the 5.0–7.0 range and for the 6.0–8.0 range.
11- Test tube
12- burner
13- Urinometer
14- Pasteur pipette
REAGENTS
12345-
20 % 5-sulfosalicylic acid
Ehrlich reagent
Acetic acid
Lougal iodine
Dipstick strip (reagent strip): should be considered as a first choice because of its
practical, easy and cost effective.
6.1.1 MACROSCOPICAL EXAMINATION OF URINE
Appearance
Urine is normally clear straw-yellow in color. More concentrated urine may appear dark
yellow, the presence of blood cells or excess salts may make the urine appear cloudy.
Pigments from bile substances may make the urine appear deep yellow or brown. Urine
can occasionally appear colorless. Report the appearance as clear or cloudy, colorless,
pale yellow, deep yellow or brown.
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Color
Urine varies in appearance, depending principally upon a body's level of hydration, as
well as other factors. Normal urine is a transparent solution ranging from colorless to
amber but is usually a pale yellow. In the urine of a healthy individual the color comes
primarily from the presence of urobilin. Urobilin in turn is a final waste product resulting
from the breakdown of heme from hemoglobin during the destruction of aging blood
cells.
Colorless urine indicates over-hydration, generally preferable to dehydration (though it
can remove essential salts from the body). Colorless urine in drug tests can suggest an
attempt to avoid detection of illicit drugs in the bloodstream through over-hydration.
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Colorless very dilute urine caused by polyuria, diabetes insipidus
Dark yellow urine is often indicative of dehydration.
Yellowing/light orange may be caused by removal of excess B vitamins from the
bloodstream.
Certain medications such as rifampin and phenazopyridine can cause orange
urine.
Reddish urine is termed hematuria, a symptom of a wide variety of medical
conditions may be caused by porphyria (not to be confused with the harmless
temporary pink or reddish tint caused by beeturia).
Dark orange to brown urine can be a symptom of jaundice, rhabdomyolysis, or
Gilbert's syndrome.
Black or dark-colored urine is referred to as melanuria and may be caused by a
melanoma.
Pinkish urine can result from the consumption of beets.
Greenish urine can result from the consumption of asparagus.
Milky caused by lipiduria
Blue urine stains can be caused by blue diaper syndrome
Odor
Normally with aromatic odor.
Abnormally:
Offensive odor due to pus increase.
Acetone odor due to ketones increase ( ketonuria).
Reaction (PH)
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Principle
Colored indicator paper is dipped in the urine (or placed in a watch glass and a few drops
of urine are added to it). The color changes according to the PH. The paper is then
compared with a standard control chart giving the corresponding pH value.
Method
The urine specimen must be tested within 1 hour of collection
1- Place a strip of universal indicator paper in a watch glass, let a few drops of fresh
urine fall from the dropper on to the paper alternatively, dip the test paper
directly into the urine in the receptacle.
2- Pick the strip of paper up with forceps. Compare the color obtained with those
shown on the standard chart .Read off the pH unit given for the color that matches
the test paper most closely.
3- According to the result obtained, select a strip of indicator paper for the
corresponding limited range. For example, if the pH is 6, use indicator paper for
the range 5.0–7.0. If the pH is 7 or more, use indicator paper for the range 6.0–
8.0. The pH of urine is normally about 6.0 (range 5.0–7.0). Acid pH values (4.5–
5.5) are observed in some forms of diabetes, muscular fatigue and acidosis.
Alkaline pH values (7.8– 8.0) are common in patients with infections of the
urinary tract and in people on a vegetarian diet. Determination of the pH of urine
is useful for the identification of crystalline
Detection of Glucose in urine
Introduction:
Glucose may appear in urine at different blood glucose level , varying in individuals
blood level, Glomerular, blood flow, tubular reabsorption rate and urine flow influence its
appearance. Glucosuria usually occur when blood level is more than 180 to 200 mg/dl
Principle
Glucose is the most commonly found sugar substance in urine, particularly in diabetic
patients and patients suffering from chronic renal failure.
method
Glucose in urine can be detected using a urine dipstick, the dipsticks are placed into the
urine and immediately removed. They are then compared with a comparison chart after
an appropriate time that is also specified on the chart
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The color changes observed on the dipstick will give a semi-quantitative estimation of the
amount of substance present. This can be reported as negative, Trace , +(160-195) , ++(
>200) , +++ (250-300) . According to the read of glucose strip or as an approximate value
of the concentration of the substance tested for. Dipsticks must be stored according to the
manufacturer’s instructions
Detection and estimation of protein
Principle:
Protein is not normally found in urine, this is because kidneys ensure that large molecules
such as protein remain in the blood. However, the kidneys sift out the smaller impurities
and expel them through the urine. In case some protein does get through, the body usually
re-absorbs it and uses it as an energy source. However, if there is too much protein in the
blood, then some amount of protein will be present in the urine. This can happen even if
the kidneys are functioning properly Elevated protein levels are observed in the urine of
patients with:

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urinary Schistosomiasis
chronic renal disease
pyelonephritis
diabetes mellitus systemic disorders (lupus erythematosus) multiple myeloma
However, orthostatic proteinuria, a form of functional proteinuria usually seen in
young men, which occurs on standing up and disappears on lying down, has no
pathological significance
METHOD
Urine for Albumin test done by one of the following: acetic acid test
1. Prepare 1 ml of urine in a test tube then heat gently till ebullition 2.If the
solution becomes turbid that indicates the presence of albumin or amorphous
urate.
So you must put acetic acid for insurance if the turbidity precipitated so it is
amorphous urate, if still turbid so it is albumin in urine.
 Sulfosalicylic acid test
1-To approximately 3ml of supernatant urine in test tube and one equal amount of
3%SSA invert in the mix
2- Let to stand exactly 10 minute
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3-Invert again twice
 Rapid test Protein can also be detected using a urine dipstick
1- Observe the degree of precipitation and grade the results according to the
following:
Table 9: Degree of Precipitation
Negative
No turbidity
Trace
perceptible
+
Distinct turbidity but no discrete
granulation
++
Turbidity with granulation but no
flocculation
+++
Turbidity with granulation and
flocculation
++++
Clumps of precipitated protein or solid
precipitate
Specific gravity
clinical significance
urinalysis is a parameter commonly used in the evaluation of kidney function and can aid
in the diagnosis of various renal diseases ( eg diabetes insipidus).
principle
Specific gravity (which is directly proportional to urine osmolality which measures solute
concentration) measures urine density, or the ability of the kidney to concentrate or dilute
the urine over that of plasma. Dipsticks are available that also measure specific gravity in
approximations. Most laboratories measure specific gravity with a refractometer Specific
gravity between 1.002 and 1.035 on a random sample should be considered normal if
kidney function is normal. Since the specific gravity of the glomerular filtrate in
Bowman's space ranges from 1.007 to 1.010, any measurement below this range indicates
hydration and any measurement above it indicates relative dehydration
Method



The Urinometer vessel is filled three fourth full with urine ( minimum volume of
urine 15 ml)
The urinometer is inserted with spinning motion
Make sure that is floating freely ( which reading the urinometer be sure that it is
not touching the sides or the bottom of cylinder avoid surface bubble read a
bottom of the meniscus
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 we can identifying the specific gravity by using dipstick
Bilirubin
Introduction
Bilirubin, a product of hemoglobin catabolism, is characterized by its distinctive yellow
pigment. The presence of bilirubin in urine is always abnormal. In most healthy
individuals the amount of conjugated bilirubin excreted is not detected by the strips. In
cases when bilirubin is elevated and is conjugated, it will be detected by the test strip. It is
important to note that unconjugated bilirubin cannot be excreted by the kidneys because it
is bound to albumin and is not soluble in water. In the liver, bilirubin combines with
glucuronic acid through the action of a glucuronyl transferase to form water soluble
bilirubin diglucuronide. Under normal circumstances, conjugated bilirubin passes from
the bile duct and then to the intestinal tract. Intestinal bacteria reduce conjugated bilirubin
to urobilinogen. Approximately half of the urobilinogen is excreted in the feces; most of
the other half is recirculated through the liver. A small amount of urobilinogen bypasses
the liver and is excreted in the urine
Normally bilirubin is not found in a routine urine test . this test is done to detect
abnormally high urine concentration of direct (conjugated ) bilirubin and help to identify
the cause of jaundice .
Clinical significance
The presence or absence of bilirubin in urine must be correlated with serum bilirubin test,
the specimen appearance is dark with yellow foam
Detection Bilirubin in urine
1. Prepare 1 ml of urine in a test tube then put 2 -3 drops Iodine on the wall of test tube
gently.
2. A violet ring appeared on the surface of urine
3. Shake the tube, then solution becomes violet or brownish
4- if violet ring appeared on the surface of urine mean positive result
urobilinogen test
Urobilinogen test is directly associated with the function of liver and any disorder or
infection related to this organ can be traced in the urobilinogen test normal specimen
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contain small amount of urobilinogen (0.1 to 1 Ehrlich units per dl of urine) this test
done to aid diagnosis of extra hepatic obstruction such as blocking of the common bile
duct and to aid differential diagnosis of hepatic and pre hepatic.
method
Urobilinogen Analysis
1234-
Add 2.5ml of afresh specimen of urine into a tube
Add o.25ml of Ehrlich reagent
Allow to stand for 5 minute
A tea color of the urine indicate urobilinogen is present in increased amounts .
A fine pink or brown color indicates that urobilinogen present in normal amount.
We can also be use the dipstick to identifying the urobilinogen in urine
The test for urobilnogen is based on the Ehrlich Aldehyde Reaction. Pdimethylaminobenzaldehyde in an acid medium with a color enhancer reacts with
urobilinogen to form a pink-red color. The urine chemical reagent strip reactivity
increases with increasing temperature. The optimum temperature for testing is 22° 26°C. A freshly voided sample is always best for ensuring optimal results
False-Positive and False-Negative Urobilinogen Results
False-positive results: A false positive urobilinogen reaction may occur with the
chemical reagent strip method when substances known to react with Ehrlich's reagent
such as sulfonamides and p-aminosalicylic acid are present in the urine. Drugs that
contain Azo dyes, such as Azo Gantrisin, have a gold color that masks the reaction,
causing a false positive reaction. Atypical color reactions may be obtained in the presence
of high concentrations of p-aminobenzoic acid. The chemical reagent strip urobilinogen
test cannot detect porphobilinogen in a urine specimen. Porphobilinogen is a molecule
formed during the synthesis of the heme portion of hemoglobin. False-negative
resultsDue to the instability of urobilinogen, a false negative result may occur using the
chemical reagent strip method if the urine specimen has remained at room temperature
for an extended period of time exposed to light. A false negative result may also occur if
formalin is present.
Detection of ketones bodies
Introduction
Normal urine does not contain ketone bodies. Acetone and other ketones bodies
may appear in urine:
 In severe or untreated diabetes( diabetic ketoacidosis )
 In certain other conditions (dehydration, vomiting, starvation and following
strenuous exercise).
Principle
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When sodium nitroprusside is added to urine containing ketone bodies, a purple color is
produced.
Materials and reagents
1- Test-tubes
2- Test-tube rack
3- Measuring cylinder, 10ml
4- Dropping pipette
5- Sodium nitroprusside crystals
6- Acetic acid
7- Ammonia.
Method
1- Just before carrying out the test, place a sufficient amount of sodium nitroprusside
crystals into a test-tube to cover the bottom
2- Add 5ml of distilled water. Shake well until the crystals are almost dissolved.
(Not all the crystals are expected to dissolve as the solution is saturated.)
3- Measure 10ml of urine into another test-tube.
4- Add four drops of acetic acid to the urine, followed by 10 drops of freshly
prepared Sodium nitroprusside solution and mix well
5- Holding the tip of the pipette against the side of the tube, let 20 drops (1 ml) of
ammonia solution flow on to the surface of the liquid Wait for 5 minutes before
reading a positive result may be obvious before this time
RESULT
If the result is positive purple ring appears on top of the urine. If the result is negative,
no color change occurs
Reporting the results of the test for detection of ketone bodies in urine
a: Positive reaction;
b: negative reaction.
substances in urine Color change Result
None = Negative Pink ring = +
Red ring= ++
Purple ring =+++
Ketone bodies in urine can also be detected using a urine dipstick
6.1.2 MICROSCOPICAL EXAMINATION OF URINE
Detection of abnormal elements
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Urine contains cells and crystals in suspension that can be collected by centrifugation or
by allowing the urine to stand and the suspended particles to form a sediment. The
resulting urinary deposit can be examined under the microscope
In certain diseases of the urinary tract, the urinary deposits are considerably altered.
Method
Preparation of the deposit
1- Mix the urine specimen gently and pour 10 ml into a centrifuge tube.
2- Centrifuge the specimen at medium speed (1500RPM) for 5 minutes.
3- Pour off the supernatant by quickly inverting the tube without shaking. (The
supernatant may be used for biochemical testing.)
4- Tap of the tube to re-suspend the deposit.
5- Transfer one drop of the deposit on to a slide using a Pasteur pipette and cover
with a cover slip
6- Label the slide with the patient’s name or identification number.
Microscopic examination
Using the X10 objective and with the condenser lowered, scan the cover slip all over to
look for ova of Schistosoma haematobium when indicated.
Using the X40 objective and with the condenser lowered or aperture reduced, scan the
coverslip area again and report any findings as a quantitative value for each high-power
field The following may be found in urine
1- Erythrocytes
2- Leukocytes
3- epithelial cells
4- casts
5- fungi
6- bacteria
7- crystals
8- parasite eggs and larvae( Schistosoma haematobium )
9- Trichomonas vaginalis
10- spermatozoa.
 Erythrocytes
Erythrocytes in urine may be:
(a) Intact: small yellowish discs, darker at the edges (8um);
(b) Crenated: spiky edges, reduced diameter (5–6um);
(c) Swollen: thin circles, increased diameter (9–10um).
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The shape of the cells often changes during storage of urine and does not have any
diagnostic importance.
In normal individual s occasional red blood cell (0-2HPF) may be seen in microscopic
examination of the sediment
Note: Erythrocytes may be found in the urine of women if the specimen has been taken
during the menstrual period.
 Leukocytes
Found in urine may be:
(a) Intact: clear granular discs, 10–15mm (the nuclei may be visible);
(b) Degenerated: distorted shape, shrunken, less granular;
(c) Pus: clumps of numerous degenerated cells.
The presence of many leukocytes, especially
in clumps, indicates a urinary tract infection.
How to express the quantity leukocytes
found in urine deposits
Place one drop of urine deposit on a slide and cover with a cover slip.
Using the x40 objective, examine the deposit and count the number of leukocytes per
microscope field.
Report the results according to cells / high power field count.
EPETHELIAL CELL
Different types of epithelial cells can be observed in urine. Some of these are readily
identifiable; however it is difficult to distinguish small transitional epithelial cells from
WBC and renal tubular epithelial cells from transitional epithelial cells. Thus, all nonsquamous cells in urine are considered to be of transitional origin. When we are unsure
about the origin of the cells in urine, we can stain a urine sediment with Wright's stain (or
Diff-quik) and perform a cytological examination on the urine sediment.
Epithelial cells are subjectively semi-quantified in urine (usually under low power using
the 10x objective) as: none seen, few, moderate, many
Transitional epithelial cells
The urinary tract from the pelvis down the ureters to the bladder and the proximal urethra
is lined by transitional epithelial cells. These cells vary in size and shape depending on
the location from which they originate, e.g. those from the renal pelvis are more caudate
whereas those from the bladder are more round to polygonal and vary in size. These cells
naturally slough into the urine in quite low numbers, so none to a few transitional
epithelial cells are seen in the urine from healthy animals. Note that this depends on the
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method of urine collection, since these cells will be sloughed (traumatically) when the
bladder is catheterized.
Transitional epithelial cells must be distinguished from WBC, because they both have the
same granular appearance. In general, transitional epithelial cells (arrow in above image)
are larger and have more irregular borders than WBC (which are uniformly more round,
arrowhead in above image).
Squamous epithelial cells
These can be keratinized or non-keratinized.
Non-keratinized squamous epithelial cells originate from the distal urethra, prepuce
and/or vagina. They are larger than transitional cells and have small central nuclei. They
can be round or have one or more flat border. Keratinized squamous epithelial cells are
from the skin or vulva and are large cells with angular borders. They may or may not
have nuclei (see upper image to the right). Nuclei are more visible in cells when the urine
is stained with Sedi-stain (see central panel on right). If in doubt about the origin of the
cells, a Wright's stain (routine hematologic stain) can be performed on a urine sediment
and demonstrates the central nuclei and angular borders of squamous epithelial cells
(lower panel on right).Squamous cells are frequently seen as contaminants in voided
urine samples and can also contaminate samples collected by catheterization
Renal tubular epithelial cells
These are rarely seen in the urine and, as mentioned above, are very difficult to
distinguish from transitional epithelial cells. If large numbers of smaller epithelial cells of
uniform appearance (size and shape) are observed in the urine, a renal origin for these
cells is suspected. Transitional epithelial cells tend to be more variable in size and shape
(to some extent). Sloughing of large numbers of renal tubular epithelial cells would
indicate renal tubular injury
RBCs
WBCs
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Squamous epithelial cells
Transitional epithelial cells
Neoplasia
Fig15: common urine cells
 CAST
Hyaline casts: These can be quite difficult to see in wet preparations of urine sediments
with light microscopy, even with the condenser of the microscope racked down. They are
much easier to visualize using phase contrast; however phase is usually not available on
most microscopes. They become more visible with regular light microscopy if fat sticks
to the protein matrix (Tamm-Horsfall mucoprotein) that makes up the hyaline cast
(hyaline with fat) or particulate material from degenerating cells is present within the cast
matrix (hyaline to finely granular cast).
Granular casts: are rather short casts filled with large granules, pale yellow in color,
with rounded ends. The granules come from degenerated epithelial cells from the tubules
of the kidney.
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Fine granular casts have smaller granules that do not fill the cast
Do not confuse with hyaline casts, partly covered by amorphous phosphate
crystals
Blood casts are filled with more or less degenerated erythrocytes, brownish in
color. They are found in acute kidney disease.
Pus casts are completely filled with leukocytes
Hyaline casts, which may contain a few leukocytes suffering from kidney
infection.
Epithelial casts are filled with pale yellow epithelial cells (To make the cells more
distinct, add a drop of 10% acetic acid to the deposit.)
Epithelial casts have no diagnostic significance.
Fatty casts are very shiny yellowish casts; the edges are indented and distinct and
the ends are rounded They are soluble in ether but not in acetic acid Fatty casts
are found in patients with severe kidney disease.
Pseudo casts Do not mistake for casts:
1- clumps of phosphate crystals, short and clear-cut
2- aggregations of translucent mucus, the ends tapering into threads
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Fig16: different casts seen in urine
Crystals
Crystals have regular geometric shapes unlike amorphous debris, which is made
up of clumps of small granules with no definite shape Except in very rare diseases,
crystals in urine have no diagnostic significance.
1- Calcium oxalate (acid urine)
 Size: 10–20µ or about 50µ
 Shape: envelope-shaped or peanut-shaped
 Color: colorless, very shiny.
2- Uric acid (acid urine)
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 Size: 30–150µ.
 Shape: varies (square, diamond-shaped, cubical or rose-shaped).
 Color: yellow or brownish-red.
3- Triple phosphates (neutral or alkaline urine)
 Size: 30–150µ.
 Shape: rectangular or like a fern leaf or star
 Color: colorless, shiny
4- Urate crystals
 Urates (alkaline urine) (Size: about 20µ.
 Shape: like a cactus or a bundle of needles
 Colour: yellow, shiny.
 Urates are often found together with phosphates.
 Calcium phosphate (neutral or alkaline urine) Size: 30–40µ.
 Shape: like a star. Colour: colourless.
5- Calcium carbonate (neutral or alkaline urine)
 Size: very small.
 Shape: similar to millet or corn grains, grouped in pairs.
 Colour: colourless.
 If acetic acid, 10% solution is added, the crystals dissolve, giving off
 bubbles of gas.
6- Calcium sulfate (acid urine)
 Size: 50–100µ.
 Shape: long prisms or flat blades, separate or in bundles.
 Calcium sulfate crystals can be distinguished from calcium phosphate crystals by
measuring the pH of the urine.
7- Amorphous phosphates (alkaline urine)
 Amorphous phosphates appear as small, whitish granules, often scattered.
8- Amorphous urates (acid urine)
Amorphous urates appear as very small, yellowish granules, which are grouped in
compact clusters. They are not soluble in acetic acid, 10% solution, but dissolve if
the urine is gently heated
(Urine kept in the refrigerator often shows a heavy precipitate of urates.)
Other crystalline deposits
The following are rarely found in the urine. When present, however, they are found
9- Cystine (acid urine)
 Size: 30–60µ.
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 Shape: hexagonal plates.
 Color: colorless, very shiny.
Cystine crystals are found only in fresh urine as they are soluble in ammonia. & they are
soluble in acetic acid, 10% solution (one drop per drop of deposit)
They are found in patients with cystin uria, a very rare hereditary disease.
10- Acetyl sulfonamides (neutral or acid urine)
 Shape: varied, but often similar to sheaves of needles.
 Acetyl sulfonamide crystals are found in the urine following treatment with
sulfonamide drugs. The presence of these crystals should be reported as they can
cause kidney damage.
Ammonium biurate
Amorphous urate
Bilirubin
Calcium carbonate
Calcium oxalate dihydrate
Magnesium ammonium
phosphate
Uric acid
Calcium oxalate
monohydrate
Calcium phosphate
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Cystine
Tyrosine
Flat plates resembling
cholesterol
Fig17: common urine crystals
Fungi


Size: 5–12µ.
Shape: round or oval bodies of various sizes found together. Do not confuse with
erythrocytes. Budding may be seen. Fungi are not soluble in acetic acid.
 Fungi are occasionally present in urine containing glucose. Check that the urine
specimen is fresh.
bacteria
In healthy persons the urine contains practically no organisms. Bacteria may be found in
patients who have an infection of some part of the urinary tract.
Urethritis, cystitis or nephritis), or where bacteria from an infection elsewhere in the body
are excreted in the urine
The urine is centrifuged at high speed and the resulting deposit is examined under the
microscope.
Parasite eggs and larvae
The following may be found:


eggs of Schistosoma haematobium: found together with erythrocytes
Trichomonas vaginalis
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Bacteria: Bacilli
Bacteria: Cocci
Schistosoma haematobium( egg)
Trichomonas vaginalis (trophozoites)
Fig18: Infectious agents
6.2 GENERAL STOOL EXAMINATION
Introduction
Many diseases are caused by infection with parasites. They are an important cause of
diarrhea (which is a major health problem in developing countries. If acute diarrhea is
caused by parasitic infection, this can be determined by examination of a stool specimen.
Clinical Significance
The Inspection of feces is important because it may lead to a diagnosis of parasitic
infestation, obstructive jaundice, diarrhea, malabsorption, rectosigmoidal ,obstruction ,
dynsentry , ulcerative colitis , or gastrointestinal tract bleeding.
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Materials and reagents:







Microscope
Microscope slides
Coverslips
Wooden applicators or wire loops (0.45mm, nickel–chromium alloy wire)
Grease pencils
Sodium chloride, 0.85% solution
Lugol iodine, 0.5% solution
Examination of stool specimens for parasites
Collection of specimens
Inspect approximately 100 g of faeces in a clean, dry container without preservatives.
A screw-top wide mouth container is most suitable. Make sure that any adult worms or
segments passed are included.
For collection of stool specimens for bacteriological examination (e.g. for culture
of cholera and other bacteria that cause dysentery),.
The container with specimen should be labeled, the urine and dirt should be excluded,
urine will destroy any amoebic trophozoites and dirt will interfere with diagnosis
 never leave the stool specimen exposed to the air in container without lead
 never keep the stool specimen artificially warm and don’t leave in the sun
Using Cary–Blair transport medium
Cary–Blair transport medium will preserve many kinds of enteric bacteria (cholera
vibrios, other vibrios, salmonella, shigella, etc.) for up to 4 weeks.
The uninoculated medium may be stored in a sealed bottle at room temperature for 8–12
weeks.
1. Dip a sterile cotton wool swab in the stool specimen
2. For infants or other patients who cannot produce a stool specimen, take a rectal swab.
Moisten the swab with sodium chloride solution and introduce the swab into the rectum.
Turn the swab several times with a circular movement
3. Place the swab in a bottle containing Cary–Blair medium (three-quarters full)
and send it to the bacteriology laboratory. If you cannot send the swab immediately,
store it at room temperature.
Important:
 Never store the swab in the incubator.
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 Never store the swab in the refrigerator.
6.2.1 Macroscopical examination
Faecal samples are best described by their colour, consistency and presence or absence of
macroscopic blood, exudate and adult parasite.
1-Colour
The color can be described as:
 Black (bleeding from upper gastrointestinal tract is more likely to cause stool
black)
 Brown
 pale yellow (fat)
 white (obstructive jaundice)
 green
 bloody (especially blood originating from lower gut may cause the stool to be
red )
 clay (suggested diminution or absence of bile or presence of barium sulfate )
2-Consistency
The consistency can be described as:
 formed (normal shape)
 soft formed
 unformed or liquid (watery).
The presence of external blood or mucus, usually seen as streaks of red or white, should
be noted. Blood may be present in certain medical conditions (e.g. ulcerative colitis,
schistosomiasis.
3- Pus
Patient with chronic ulcerative colitis and chronic bacillary dysentery frequently pass
large quantity of pus with the stool
 Large amount of pus never accompany amoebic colitis .
 No pus seen in watery stool ….. viral gastroenteritis.
4- Mucus
Even in slightest quantity is abnormal
Bloody mucus = inflammation of rectal canal
Associated with pus and blood is found in stool of patient with ulcerative colitis ,
bacillary dysentery , intestinal tuberculosis
Mucus with pus and blood = ulcerative colitis , bacillary dysentery . ulcerative carcinoma
of colon . intestinal tuberculosis
5 - blood
Blood in stool should be never ignored
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 Bleeding upper GIT may give black- tarry appearance to stool.
 Bleeding from lower GIT may give red color
6- Fecal fat
 It is raised in malabsorption.
6.2.2 Microscopic examination
Direct microscopic examination of faeces in saline or iodine suspension is useful for the
following reasons:
 to detect motile trophozoites
 to detect ova and cysts present in moderate numbers
 to detect erythrocytes, cellular debris or excess fat
Select unformed or liquid faeces when using direct microscopy for detection of
trophozoites. Formed stools rarely contain motile trophozoites. Also perform a direct
examination of any external blood or mucus.
Method
1. Prepare a 1:1 mixture of 0.5% Lugol iodine solution.
2. Take a dry microscope slide and label it with the name or number of the patient.
3. Put: one drop of sodium chloride solution warmed to 37°C in the middle of the
left half of the slide; and one drop of the iodine–acetic acid solution in the middle
of the right half of the slide
4. Using an applicator or wire loop, take a small portion (about 2–3mm diameter) of
the stool.
a) If the stools are formed, take the portion from the centre of the sample and
from the surface to look for parasite eggs.
b) If the stools contain mucus or are liquid, take the portion from the mucus
on the surface or from the surface of the liquid to look for amoebae.
5. Mix the sample with the drop of sodium chloride solution on the slide.
6. Using the applicator or wire loop, take a second portion of stool from the
specimen as described above and mix it with the drop of iodine, acetic acid
solution, discard the applicator (or flame the wire loop) after use.
7. Place a coverslip over each drop (apply the coverslips as shown in to avoid the
formation of air bubbles).
8. Examine the preparations under the microscope. For the saline preparation use the
x10 and x40 objectives and a x10 eyepiece. As the eggs and cysts are colorless,
reduce the amount of light using the condenser aperture or lower the condenser to
increase the contrast. Examine the first preparation with the x10 objective, starting
at the top left hand corner as indicated in Focus on the edge of one coverslip using
173 | P a g e
the x10 objective and examine the whole area under each coverslip for the
presence of ova and larvae of parasit . Then switch to the x40 objective and again
examine the whole area of the coverslip over the saline for motile trophozoites
and the area of the coverslip over the iodine for cysts.
9. Lugol iodine–acetic acid solution causes the trophozoite forms to become
nonmotile The nucleus is clearly stained but it may be difficult to distinguish
between trophozoite and cystic forms.
10. Using a fine Pasteur pipette, allow a drop of 5% methylene blue solution to run
under the coverslip over the saline preparation This will stain the nuclei of any
cells present and distinguish the lobed nuclei of polymorphs from the large single
nuclei of mucosal cells.
Intestinal protozoa
Protozoa are microorganisms consisting of a single cell. Intestinal protozoa may be
found in stools in their motile form (trophozoites) or as cysts. Some intestinal protozoa
are pathogenic (others are harmless. All these protozoa are found throughout the world.
Identification of motile forms (trophozoites)
The trophozoites of protozoa are motile
 either because of slow movements of the cell (amoebae)
 or because they have rapidly moving flagella (long whip-like threads) or
cilia (numerous short hairs).
Trophozoites are chiefly found in:
 watery stools
 stools containing mucus
 soft formed stools.
The following features are useful for the identification of motile forms of intestinal
protozoa










size
cytoplasm
pseudopodia
nuclei
Ectoplasm
Endoplasm
Vacuoles
Inclusion bodies containing erythrocytes, bacteria, yeast cells, debris, etc.
nuclear membrane (chromatin)
nuclear karyosome
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 flagella
 Undulating membrane.
Identification of cysts
Cysts are the resistant forms of certain intestinal amoebae, flagellates and ciliates.
They are small, round and non-motile and may have one or several nuclei.
Measurement of cysts is useful for the correct identification of species.
Importance of cysts
The clinical importance of cysts varies from country to country. The cyst is the infective
form of the organism. Healthy persons may be asymptomatic carriers of cysts and are
therefore, a public health hazard.
The most important problem in the laboratory is the precise identification of cysts
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Taenia saginata
Hymenolepis nana
Diphyllobothrium latum
Ascaris lumbricoides
Ancylostoma duodenale
Trichuris trichuria
Strongyloides stercoralis
Clonorchis sinensis
Paragonimus westermani
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Schistosome mansoni ova
Schistosome japonicum
Schistosoma haematobium
Figure19: Ova in GSE
Entamoeba histolytica cyst
Entamoeba histolytica trophozoites
Giardia lamblia cyst
Giardia lamblia trophozoites
Figure 20: Entamoeba histolytica & Giardia lamblia (HPF)
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Module 4
Session 2: Urine culture, throat swab, and nasal swab
Objectives
At the end of this session participants will be able to:
1. Explain urine culture
2. Explain throat swab
3. Explain nasal swab
Trainers Preparation:




Review the reading material and the session plan.
Prepare the presentation as appropriate and as recommended in the method
column of the session plan, or write the information on a flipchart or board where
all participants can see it.
Prepare copies of the reference materials/handouts and exercises.
Arrange the training room.
Methods and activities

Discussion-lecture, question & answers
Evaluation/assessment
Questions and answers, trainer’s observation
Estimated Time
165 minutes
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Session plan -2Objectives
4.2.1. Explain urine culture
Content





4.2.2. Explain throat swab




4.2.3. Explain nasal swab



Methodology
Introduction
Media and
diagnostic reagents
Specimen collection
Screening method
Calibration
Discussion -lecture
Introduction
Media and
diagnostic reagents
Bacterial agents of
pharyngitis
Culture and
identification
Discussion -lecture
60 minutes
60 minutes
Question & answers
Specimen
Collection,
Transport and
Storage
Correct specimen
type and method of
collection
Staphylococci
Identification
45 minutes
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6.3 Urine culture
Introduction
Urine is the specimen most frequently submitted for culture. It also presents major
problems in terms of proper specimen collection, transport, culture techniques, and
interpretation of results. As with any other specimen submitted to the laboratory, the
more information provided by the submitting physician the more able is the laboratory to
provide the best possible culture data.
The most common sites of urinary tract infection (UTI) are the urinary bladder (cystitis)
and the urethra. From these sites the infection may ascend into the ureters (ureteritis) and
subsequently involve the kidney (pyelonephritis). Females are more prone to infection of
the urinary tract and also present the greater problem in the proper collection of
specimens.
In both males and females, UTI may be asymptomatic, acute, or chronic. Asymptomatic
infection can be diagnosed by culture. Acute UTI is more frequently seen in females of
all ages; these patients are usually treated on an outpatient basis and are rarely admitted to
hospital. Chronic UTI in both males and females of all ages is usually associated with an
underlying disease (e.g., pyelonephritis, prostatic disease, or congenital anomaly of the
genitourinary tract) and these patients are most often hospitalized. Asymptomatic, acute,
and chronic UTI are three distinct entities and the laboratory results often require
different interpretation.
Asymptomatic pyelonephritis in females may go undetected for some time, and is often
only diagnosed by carefully performed quantitative urine culture. Chronic prostatitis is
common and difficult to cure, and is often responsible for recurring UTI. In most UTI,
irrespective of type, enteric bacteria are the etiological agents, Escherichia coli being
isolated far more frequently than any other organism. In about 10% of patients with UTI,
two organisms may be present and both may contribute to the disease process. The
presence of three or more different organisms in a urine culture is strong presumptive
evidence of improper collection or handling of the urine specimen. However, multiple
organisms are often seen in UTI in patients with indwelling bladder catheters
Expected pathogens
- Enterococci
- Escherichia coli
- Mycobacterium tuberculosis
- Other Enterobacteriaceae
- Other staphylococci
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- Pseudomonas and other non-fermenters
- Staphylococcus saprophyticus
Media and diagnostic reagents
- Isolation and quantitative media
Blood agar
CLED agar or chromogenic media
MacConkey agar, with crystal violet, )
- Identification media and diagnostic reagents
Kligler iron agar (KIA)
Kovacs reagent for indole
motility–indole–urease (MIU) medium
oxidase reagent
lysine–decarboxylase broth (Möller)
ONPG test
Simmons citrate agar
For staphylococci and enterococci
catalase test (H2O2)
coagulase plasma
bile–aesculin agar (for enterococci)
novobiocin (5mg) disc for differentiating negative-coagulase staphylococci
Specimen collection
It cannot be overemphasized that considerable importance is attached to the method of
collection of urine specimens, transport to the laboratory, and the initial efforts by the
laboratory to screen and culture the urine. It is the responsibility of the laboratory to
provide the physician with sterile, wide-mouthed, glass or plastic jars, beakers, or other
suitable receptacles. Urine specimens may have been collected by a surgical procedure,
e.g., suprapubic aspiration, cystoscopy, or catheterization. If not, the laboratory must
insist on a clean-catch midstream urine specimen, particularly in females and children.
Since urine itself is a good culture medium, all specimens should be processed by the
laboratory within 2 hours of collection, or be kept refrigerated at 4 °C until delivery to the
laboratory and processed no longer than 18 hours after collection.
Whenever possible, urine specimens for culture should be collected in the morning. It is
advisable to ask the patient the night before to refrain from urinating until the specimen is
collected.
Women
A woman who is ambulatory should:
1. Wash her hands thoroughly with soap and water and dry them with a clean towel.
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2. Undress in a suitable room, spread the labia, and cleanse the vulva and labia
thoroughly, using sterile cotton gauze pads and warm soapy water wiping from front to
rear.
3. Rinse thoroughly with warm water and dry with a sterile gauze pad. During the entire
process the patient should keep the labia separated and not touch the cleansed area with
the fingers.
4. Pass urine, discarding the first part of the stream. Collect the remaining urine in the
sterile container, closing the lid as soon as the urine has been collected.
5. Hand the clean midstream urine, in the closed container, to the nursing personnel for
prompt delivery to the laboratory.
For bedridden patients, the same procedure is followed, except that a nurse must assist
the patient or, if necessary, do the entire cleansing procedure before requesting the patient
to pass urine.
In both situations every effort must be made to collect a clean-catch urine specimen in a
sterile container and to ensure that it is delivered promptly to the laboratory together with
information on the patient, clinical diagnosis, and the requested procedures.
Men
A man who is ambulatory should:
1. Wash his hands.
2. Pull back the foreskin (if not circumcised) and pass a small amount of urine.
3. Still holding back the foreskin, pass most of the remaining urine into a sterile
container. This is a midstream urine specimen.
4. Place the cover on the container and hand to nursing staff for prompt delivery to the
laboratory.
For bedridden patients:
1. If necessary, nursing personnel should pull back the foreskin, wash and dry the glans
with soapy water and gauze pads.
2. With foreskin pulled back, the patient should pass a small amount of urine into a
urinal.
3. The patient should then pass most of the remaining urine into the sterile container. The
cover should be placed on the container and the specimen transported to the laboratory.
Infants and children
Collection of a clean-catch urine specimen from infants and children who are ill in bed or
uncooperative can be a problem. Give the child water or other liquid to drink. Clean the
external genitalia. The child can be seated on the lap of the mother, nurse, or ward
attendant, who should then encourage the child to urinate and collect as much urine as
possible in a sterile container. The container should then be covered and delivered to the
laboratory for prompt processing.
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Culture and interpretation
All urine specimens brought to the microbiology laboratory should be examined at once,
or placed in a refrigerator at 4 °C until they can be examined. The examination procedure
includes the following steps:
1. Examination of a Gram-stained smear.
2. A screening test for significant bacteriuria.
3. A definitive culture for urine specimens found to be positive in the screening test (step
2), and for all specimens obtained by cystoscopy, Suprapubic aspirate (SPA) , or
catheterization.
4. Susceptibility tests on clinically significant bacterial isolates.
Preparation and examination of a Gram-stained smear is a necessary part of the
laboratory process. Using a sterile Pasteur pipette (one for each sample), place one drop
of well mixed, uncentrifuged urine on a slide. Allow the drop to dry without spreading,
heat-fix and stain. Examine under an oil-immersion lens for the presence or absence of
bacteria, polymorphonuclear leukocytes, and squamous epithelial cells.
One or more bacterial cells per oil-immersion field usually implies that there are 105 or
more bacteria per milliliter in the specimen. The presence of one or more leukocytes per
oil-immersion field is a further indication of UTI. Non-infected urine samples will
usually show few or no bacteria or leukocytes in the entire preparation. In specimens
from females, the presence of many squamous epithelial cells, with or without a mixture
of bacteria, is strong presumptive evidence that the specimen is contaminated with
vaginal flora and a repeat specimen is necessary, regardless of the number of bacteria per
oil-immersion field. If results are required urgently, the report of the Gram-stain findings
should be sent to the physician with a note that the culture report is to follow.
Screening method
The absence of leukocytes and bacteria in a Gram-stained smear of a clean-catch urine
sample prepared as described above is good evidence that the urine is not infected. A
urine specimen that is “negative” on careful examination of the Gram-stained smear does
not need to be cultured. An alternative simple and effective screening test is the test strip
for leukocyte esterase/nitrate reduction. The strip is dipped into the urine specimen as
instructed in the package literature. Any pink colour is a positive reaction indicating the
presence of leukocyte esterase and/or bacteria in excess of 105 per ml. Urine samples that
are positive in the screening test should be cultured as soon as possible to prevent
possible overgrowth by nonsignificant bacteria. If the strip does not develop a pink colour
it is interpreted as a negative screening test, is so reported, and no culture is indicated.
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The test strip may not be sensitive enough to detect bacterial counts of less than 105 per
ml of urine.
Quantitative culture and presumptive identification
Calibrated loop technique
There are several culture methods for the quantification of bacteria in urine. The easiest
and most commonly used are the calibrated loop technique. Of these method using CLED
or chromogenic media, are considered to be the most versatile and efficient for large
numbers of specimens.
Chromogenic media contain various substrates which permit presumptive identification
of several common species through a change in either colony pigmentation or colour of
agar. They perform satisfactorily compared to CLED and have the advantage that mixed
cultures are easier to detect. However, chromogenic media from different manufacturers
can vary in specificity, and are relatively expensive Calibrated loop/surface streak
method
Mix the urine gently to avoid foaming.
Dip the end of a sterile calibrated loop (eg 1 μL, 2 μL or 10 μL) in the urine to just below
the surface and remove vertically, taking care not to carry over any on the shank.
Use this to inoculate CLED or chromogenic agar plate and spread according to the
number of specimens . A maximum of four samples per 9 cm plate is recommended for
this method with a 1 μL or 2 μL loop, or two samples if using a 10 μL loop. If a 1 μL
loop is used, one colony equals 1000 cfu/mL (ie 1 x 106 cfu/L).
SPA, other surgically obtained urine, and urine samples with expected significant
bacteriuria as low as 105 cfu/L (increased inoculum sizes are required) ,Inoculate 100 μL
(0.1 mL) of specimen aseptically to a full CLED or chromogenic agar plate.
Spread inoculum over entire surface of plate with a sterile loop or a spreader. Do not use
a sterile swab which will absorb much of the inoculum. To isolate individual colonies,
spread inoculum with a sterile loop.
No. of cfu/L = No. of cfu on plate x 104.
This semi quantitative method is only sensitive for screening down to106 cfu/L if a 5 μL
or 10 μL loop is used (eg 5 or 10 colonies), or107 cfu/L if a 1 μL or 2 μL loop is used (eg
10 or 20 colonies). (See table below).
Guidance on assessing colony counts :
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Interpretation of culture
The basis studies for interpreting urine culture results showing that bacterial counts of
≥108 cfu/L (≥105 cfu/mL) are indicative of an infection and counts below this usually
indicate contamination.
In specific patient groups , counts between 108 cfu/L (105 cfu/mL) and 105 cfu/L (102
cfu/mL) may be significant. A pure isolate with counts between 107 and 108 cfu/L (104105 cfu/mL) should be evaluated based on clinical information or confirmed by repeat
culture. Overall the confirmation of a UTI requires the demonstration of significant
bacteriuria by quantitative culture (defined according to patient group or specimen type).
Routine culture methods may not be sensitive enough to detect low bacteria levels (eg
≥107 cfu/L / ≥104 cfu/mL) and increased sensitivity will be achieved by increasing the
inoculum size .
The following is a list of imperial units and their equivalents.
≥105 cfu/mL equivalent to ≥108 cfu/L
<105 cfu/mL equivalent to <108 cfu/L
104 cfu/mL equivalent to107 cfu/L
<104 cfu/mL equivalent to <107 cfu/L
103 cfu/mL equivalent to 106 cfu/L
<103 cfu/mL equivalent to <106 cfu/L
102 cfu/mL equivalent to 105 cfu/L
Increased inoculum sizes are also required for persistently symptomatic patients without
bacteriuria if the patient has recurrent ‘‘sterile pyuria’’, or for specimens where lower
counts are to be expected, such as SPA or other surgically obtained urine.
Identification
Identification should be performed as rapidly as possible. Isolation of uropathogens
should be performed by a surface streak procedure on both blood and MacConkey agar
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using calibrated loops for semi-quantitative method and incubated aerobically at 37 °C
for 24 hours, and those cultures which becomes negative at the end of 24 hrs incubations
will be further incubated for 48 hours. Bacterial identification should be made using
biochemical tests, namely indole, citrate, oxidase, H2S production, lysine decarboxylase,
lactose fermentation, urea hydrolysis, gas production, catalase, coagulase, manitol
fermentation and novobiocin susceptibility test (See Flowcharts below ).
Clinical specimen
Primary isolation plate
(blood, CLED, Staph/Strep medium or fastidious anaerobe agar)
Gram stain
Gram positive cocci in pairs and/or short chains
Positive
(Probable Staphylococcus)
A weak catalase or pseudocatalase reaction may be
produced by some strains of Aerococcus &
Enterococcus species
Catalase
Negative
non-haemolytic
a-haemolytic
(Consider Gemella)
b-haemolysis
(Consider Leuconostoc,
Gemella, Helcococcus)
Suspected Enterococcus
(1-2 mm may be a, β or non-haemolytic.
Consider clinical details)
Optochin
Sensitive
Lancefield Group
Resistant
Rapid Aesculin hydrolysis
(or PYR or group)
A,B,C,D,F,G
Positive
S. pneumoniae:
Some S. pneumoniae
may be resistant to
optochin: if there is a
clinical suspicion of
pneumococcal infection,
confirm by performing
bile solubility
Non groupable
(A,C,G, consider
S.anginosus
group)
Negative
(repeat, consider Listeria,
check previous
tests)
“viridans”
Streptococci:
Occasional strains of
S. oralis may be
optochin sensitive:
S. pseudopneumoniae
optochin sensitive, bile
solubility
inconclusive
Enterococcus spp
(PYR & LAP positive)
PYR positive, LAP positive
consider Aerococcus, Lactococcus,
Facklamia, Dolosigranulum.
PYR positive, LAP negative
consider Globicatella, Aerococcus
viridans, Dolosicoccus
Consider
Aerococcus,
Pediococcus,
Lancefield Group B
S. anginosus group
(Aesc+, PYR-)
PYR
Negative
Positive
S. anginosus group
Group A
Further identification if clinically indicated
Commercial identification system or other biochemical identification or send to the
Reference Laboratory
Fig 21: Identification of Streptococci and Enterococci Flowchart
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Clinical specimens
Primary isolation plate
Opaque, white, cream, yellow or orange colonies on blood agar
Gram stain
Gram positive cocci in clusters
If there is a different Gram stain
appearance refer to the appropriate
SMI
Negative
(S. anaerobius is catalase
negative)
Consider other
organisms
Modified oxidase
Resistant
Positive
Micrococcus
(Aerobic growth only)
Suspected
S. aureus
Catalase Positive
Novobiocin
Negative
Sensitive
DNase, clumping factor
(slide or commercial
latex kit) Protein A or
thermostable nuclease
Negative
S. sciuri Group
(Aesculin Positive)
(Meticillin resistant strains may be
mistaken for MRSA and will grow
well on many chromogenic
selective agars).
S. saprophyticus
Group
Coagulase-negative
Staphylococcus
Positive
Confirm with tube
coagulase if required
S. aureus
Further identification
if clinically indicated
Commercial identification
system
Confirm with tube
coagulase if required
DNA +ve: S. aureus, S. intermedius, S. schleiferi, S. hycius, S. sciuri, S. chromogenes
Protein A latex +ve: S.aureus (S. saprophyticus and S. sciuri may give false positive)
Coagulase +ve: S. aureus, S. intermedius, S. hyicus, S. schleiferi
Fig 22: Presumptive Identification of Staphylococcus Species Flowchart
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Clinical Specimens
Primary isolation plate
BA
CLED B or CLED A, MAC
DCA, XLD, CT-SMAC, TCBS, CIN agar
Carbohydrate
fermenting
Carbohydrate
non fermenting
Further identification if
clinically indicated
Oxidase
Performed from non selective
medium
Negative
Further identification
Serology for possible:
Salmonella / Shigella species (XLD / DCA)
E. coli O157 (CT-SMAC) all presumptive (locally confirmed)
E. coli O157 should be sent to the Reference Laboratory Y. enterocolitica (CIN)
(see ID 20, 21, 22, 24)
Commercial identification system
or
other biochemical identification
or
send to the Reference Laboratory
Positive
Possible Pseudomonas species
or
Pasteurella species
(see ID 17 & 13)
Fig 23: Identification of Enterobacteriaceae Flowchart
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Clinical specimens
Primary isolation plate
Pseudomonas selective agar
Blood agar
(or other primary isolation media)
Burkholderia cepacia
selective agar
Typical colonies of GNR at 16-48 hr (see Sections 3.2 and 3.3)
Burkholderia cepacia complex may require up to 5 days incubation
Pigment production
Blue-green colonies
No blue-green colonies
Pink colonies on B. cepacia
selective agar
Oxidase test
Oxidase test
Gram stain of
pure culture
Positive
P. aeruginosa
growth at
42°C
Negative
Positive
Negative
Gram
negative rod
Further identification if clinically indicated
Commercial identification kit or other biochemical
identification or send to the Reference Laboratory
If required, save the pure isolate on an agar slope
Other
Discard
Fig 24: Identification of Glucose Non -Fermenting Gram negative Rods
Flowchart
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Susceptibility tests
Susceptibility tests should only be performed on well isolated colonies of similar
appearance that are considered significant according to the guidelines presented above.
Susceptibility tests are generally more important on cultures obtained from patients who
are hospitalized or have a history of recurring UTI. Cultures from patients seen in the
clinic with a primary UTI may not require a susceptibility test.
6.4 Throat swab
The upper respiratory tract can be the site of several types of infection:
 pharyngitis, sometimes involving tonsillitis, and giving rise to a “sore throat”
 nasopharyngitis
 otitis media
 Sinusitis
 epiglottitis.
Of all those infections, pharyngitis is by far the most frequent; in addition, the untreated
infection may have serious sequelae. Only pharyngitis will be considered here.
Bacteriological diagnosis of pharyngitis is complicated by the fact that the oropharynx
contains a heavy, mixed, normal flora of aerobic and anaerobic bacteria. The normal flora
generally outnumbers the pathogens and the role of the bacteriologist is to distinguish
between the commensals and the pathogens. Where possible only the latter should be
reported to the physician.
Normal flora of the pharynx
The normal flora of the pharynx includes a large number of species that should be neither
fully identified nor reported when observed in throat cultures:
• viridans (a-haemolytic) streptococci and pneumococci
• Nonpathogenic Neisseria spp.
• Moraxella (formerly Branhamella) catarrhalis (this can also be a respiratory pathogen)
• Staphylococci (S. aureus, S. epidermidis)
• diphtheroids (with the exception of C. diphtheriae)
• Haemophilus spp.
• Yeasts (Candida spp.) in limited quantity
• Various strictly anaerobic Gram-positive cocci and Gram-negative rods, spirochaetes
and filamentous forms.
The throats of elderly, immunodeficient, or malnourished patients, particularly when they
have received antibiotics, may be colonized by Enterobacteriaceae (Escherichia coli,
Klebsiella spp., etc.) and by the nonfermentative Gram-negative groups (Acinetobacter
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spp. and Pseudomonas spp.). Such patients may also have in their pharynx a proliferation
of S. aureus or of
Candida spp., or other yeast-like fungi. Although these microorganisms do not cause
pharyngitis, except in association with granulocytopenia, it is advisable to report such
isolates to the clinician, as they occasionally indicate the existence of (or may sometimes
give rise to) a lower respiratory tract infection
(e.g. pneumonia) or bacteraemia. However, an antibiogram should not be performed
routinely on these colonizing microorganisms.
Expected pathogens
 Candida albicans (oropharynx)
 Corynebacterium diphtheriae (throat and nose)
 Haemophilus influenzae (ear and sinus)
 Moraxella catarrhalis (ear and sinus)
 Neisseria meningitides
 Pseudomonas
 Staphyloccoccus aureus (ear and sinus)
 Streptococcus pneumoniae (ear and sinus)
 Streptococcus pyogenes (group A, throat)
Media and diagnostic reagents
Isolation media
Isolation media
Blood agar (prepared from a glucose-free base)
Chocolate agar
Löffler coagulated serum or Dorset egg medium
Tellurite blood agar
Modified Thayer–Martin medium (for gonococci andmeningococci)
Priority
grading
1
2
2
2
3
Diagnostic reagents
Diagnostic reagents
Bacitracin disc
Catalase and coagulase reagents
Optochin disc
Carbohydrate degradation media for Neisseria spp.
Oxidase reagent
V and XV factors (discs or strips)
Tributyrin
Priority
grading
1
1
1
2
2
2
3
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Rapid diagnostic tests
Grouping kit for haemolytic streptococci
3
Bacterial agents of pharyngitis
Streptococcus pyogenes (Lancefield group A) is by far the most frequent cause of
bacterial pharyngitis and tonsillitis. This infection is particularly prevalent in young
children (5–12 years).
Non-group-A, b-haemolytic streptococci (e.g. groups B, C and G) are uncommon causes
of bacterial pharyngitis and if detected should be reported.
Corynebacterium diphtheriae is the cause of diphtheria, a disease that is endemic in many
countries. It can reach epidemic proportions in countries where the vaccination
programme has been interrupted. Characteristically (with a few exceptions), C.
diphtheriae causes a typical form of infection, characterized by a greyish-white
membrane at the site of infection (pharynx, tonsils, nose, or larynx). Diphtheria is a
serious disease and the diagnosis is made on the basis of clinical findings. The physician
would then generally make a specific request to culture for diphtheria bacilli.
(Vincent angina) is a rare condition characterized by a necrotic ulceration of the pharynx
with or without formation of a pseudomembrane. It is associated, at the site of infection,
with a heavy mixed flora of strict anaerobes dominated by Gram negative fusiform rods
and spirochaetes, generally referred to as Fusobacterium spp. and Treponema vincentii,
and possibly others. Although both species belong to the normal mouth flora, their
presence in large numbers in a Gram-stained smear of ulcerated lesions should be
reported as a “fusospirochaetal complex”. This microscopic diagnosis need not be
confirmed by anaerobic culture, which is difficult and time-consuming. However, the
presence of this complex does not exclude the need to search for other pathogens,
particularly S. pyogenes.
Although small numbers of C. albicans or other Candida species may be part of the
normal oral flora, oral candidiasis results when the number of organisms increases
considerably in certain pathological conditions, e.g. in malnourished premature babies, in
immunodeficient adults (e.g. patients with HIV/AIDS), or in patients who have received
broad-spectrum antimicrobials or cancer therapy. The affected area—tongue, tonsils,
throat or buccal mucosa —may be extremely red, or covered with white patches or a
confluent grey-white membrane (thrush). The diagnosis of candidiasis is best made by
finding numerous yeast cells, some of them forming long mycelium-like filaments, in a
Gram-stained smear of the exudate. Swabs from the upper respiratory tract may be
submitted to the laboratory, not for the diagnosis of a clinical infection, but to detect a
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potential pathogen in a healthy subject, a pharyngeal or a nasal “carrier”. This should
only be done as part of well-defined epidemiological surveys.
The following pathogens can give rise to a carrier state in the upper respiratory tract:
• Staphylococcus aureus. Sampling of patients and staff for nasal carriers is sometimes
performed as part of an investigation of hospital outbreaks of Meticillin-resistant S.
aureus (MRSA).
• Neisseria meningitidis. Carriage of meningococci may be very prevalent (20% or more)
even at non-epidemic times. Identification of pharyngeal carriers of meningococci is
rarely needed, and need not be performed prior to the administration of prophylactic
antibiotics to family or other close contacts of patients with meningococcal disease.
• Streptococcus pyogenes. Carriage of this organism in low numbers may be prevalent,
especially among schoolchildren (20–30%).
• Corynebacterium diphtheriae. The carrier rate of the diphtheria bacillus is high in nonvaccinated populations. In such communities, it may be justified to identify and treat
carriers among the close contacts of a patient with proven diphtheria. Carriers are rare
when an immunization programme is correctly implemented.
Collection and dispatch of specimens
Ideally, specimens should be collected by a physician or other trained personnel.
The patient should sit facing a light source. While the tongue is kept down with a tongue
depressor, a sterile cotton-wool swab is rubbed vigorously over each tonsil, over the back
wall of the pharynx, and over any other inflamed area. Care should be taken not to touch
the tongue or buccal surfaces.
It is preferable to take two swabs from the same areas. One can be used to prepare a
smear, while the other is placed into a glass or plastic container and sent to the laboratory.
Alternatively, both swabs may be placed in the container and dispatched to the
laboratory. If the specimen cannot be processed within 4 hours, the swab should be
placed in a transport medium (e.g. Amies or Stuart).
Direct microscopy
The fusospirochaetal complex of necrotizing ulcerative pharyngitis (Vincent angina) and
Candida are best recognized on a Gram-stained smear, which should be prepared if the
physician makes a special request. The Gramstained smear is not useful for the detection
of streptococci or Neisseria spp.
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Moreover, the direct smear has poor sensitivity and specificity for the detection of the
diphtheria bacillus, unless the specimen has been collected with care and is examined by
an experienced microbiologist. In the absence of a physician’s request or of clinical
information, a Gram-stained smear should not be made for throat swabs.
Culture and identification
Culture for Streptococcus pyogenes
Immediately upon receipt in the laboratory, the swab should be rubbed over one-quarter
of a blood agar plate, and the rest of the plate streaked with a sterile wire loop. The blood
agar should be prepared from a basal agar medium without glucose (or with a low
glucose content), e.g. tryptic soy agar (TSA). Acidification of glucose by S. pyogenes
inhibits the production of haemolysin. Blood from any species, even human blood (fresh
donor blood),
can be used at a concentration of 5%. The plates should be filled to a depth of 4–5 mm.
Sheep blood is preferred because it does indicate haemolysis of some commensal
Haemophilus spp. and it gives no haemolysis with the zymogenes variant of
Enterococcus faecalis.
The recognition of b-haemolytic colonies can be improved, and their presumptive
identification hastened, by placing a co-trimoxazole disc (as used for the susceptibility
test) and a special low-concentration bacitracin disc over the initial streaked area.
Because S. pyogenes is resistant and many other bacteria are susceptible to cotrimoxazole, this disc improves the visibility of b-haemolysis. Incubation in a candle-jar
will detect most b-haemolytic streptococci.
A simple way to increase haemolysis is to stab the agar surface perpendicularly by
inserting the loop deep into the medium to encourage growth
of subsurface colonies. After 18 hours and again after 48 hours of incubation at 35–37
∞C, the blood plates should be examined for the presence of small (0.5–2 mm) colonies
surrounded by a relatively wide zone of clear haemolysis.
After Gram-staining to verify that they are Gram-positive cocci, the colonies should be
submitted to specific identification tests for S. pyogenes. For clinical purposes,
presumptive identification of S. pyogenes is based on its susceptibility
to a low concentration of bacitracin. For this purpose, a special differential disc is used
containing 0.02–0.05 IU of bacitracin. The ordinary discs used in the susceptibility test,
with a content of 10 units, are not suitable for identification. A b-haemolytic
streptococcus showing any zone of inhibition around the disc should be reported as S.
pyogenes. If the haemolytic colonies
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are sufficiently numerous, the presence or absence of an inhibition zone may be read
directly from the primary blood agar plate. If the colonies are less numerous, one or two
should be picked from the primary plate, streaked on one-fifth of another plate to obtain
confluent growth, and each inoculated area covered with a bacitracin disc. After
overnight incubation, the subcultures should be read for inhibition zones.
In some laboratories this presumptive identification is confirmed by serological
demonstration of the specific cell wall polysaccharides. This can be done either by the
classical precipitin method, or more rapidly by using a commercial
kit for the rapid slide coagglutination or latex agglutination tests. If desirable, bacitracinresistant b-haemolytic streptococci can be further identified using some simple
physiological tests (see Table 19). Minute colonies of b-haemolytic streptococci may be
encountered, which, when grown and serologically grouped, react with group A
antiserum. These streptococci are not
considered to be S. pyogenes and are not associated with the serious infections caused by
group A streptococci.
In reporting the presence of S. pyogenes in a throat culture, a semiquantitative answer
should be given (rare, +, + +, or + + +). Patients with streptococcal pharyngitis generally
show massive growth of S. pyogenes, with colonies over the entire surface of the plate.
Plates of carriers generally show fewer than 20 colonies per plate. Even rare colonies of
b-haemolytic streptococci should be confirmed and reported.
Table 10: Differentiation of b-haemolytic streptococci
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Culture for Corynebacterium diphtheriae
Although the diphtheria bacillus grows well on ordinary blood agar, growth is improved
by inoculating one or two special media:
• Löffler coagulated serum or Dorset egg medium. Although not selective, both of these
media give abundant growth of the diphtheria bacillus after overnight incubation.
Moreover, the cellular morphology of the bacilli is more “typical”: irregularly stained,
short to long, slightly curved rods, showing metachromatic granules, and arranged in a V
form or in parallel
palisades. Metachromatic granules are more apparent after staining with methylene blue
or Albert stain than with the Gram stain.
• A selective tellurite blood agar. This medium facilitates isolation when the bacilli are
few in number, as is the case for healthy carriers. On this medium, colonies of the
diphtheria bacillus are greyish to black and are fully developed only after 48 hours.
Suspicious colonies, consisting of bacilli with a coryneform morphology on the Gramstained smear, should
be subcultured to a blood agar plate to check for purity and for “typical” morphology. It
should also be remembered that colonies of the mitis biotype of C. diphtheriae, which is
the most prevalent, show a marked zone of b-haemolysis on blood agar.
A presumptive report on the presence of C. diphtheriae can often be given at this stage.
However, this should be confirmed or ruled out by some simple biochemical tests and by
demonstration of the toxigenicity. As the latter requires inoculation of guinea-pigs or an
in vitro toxigenic test (Elek) and has to be performed in a central laboratory, only rapid
biochemical identification will be covered here. C. diphtheriae is catalase- and nitratepositive. Urea is not hydrolysed. Acid without gas is produced from glucose and maltose,
generally
not from saccharose. The fermentation of glucose can be tested on Kligler medium.
Urease activity can be demonstrated on MIU and nitrate reduction in nitrate broth in the
same way as for Enterobacteriaceae. For the fermentation of maltose and saccharose,
Andrade peptone water can be used as a base
with a 1% final concentration of each carbohydrate. Results can usually be read after 24
hours, although it may be necessary to reincubate for one night.
It must be emphasized that the microbiology laboratory’s role is to confirm the clinical
diagnosis of diphtheria. Therapy should not be withheld pending receipt of laboratory
reports.
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Susceptibility testing
Routine susceptibility tests on throat or pharyngeal isolates are most often not required,
and may even be misleading. The major pathogens involved in bacterial pharyngitis are
S. pyogenes and C. diphtheriae benzylpenicillin and erythromycin are considered as the
antimicrobials of choice to treat both types of infection. In cases of diphtheria, treatment
with antitoxin is also indicated.
6.5 Nasal Swab
Nasal colonization with Staphylococcus aureus increases the risk of staphylococcal
infections at other sites of the body such as postoperative wounds and dialysis access
sites3. It is also associated with recurrent skin infections and nosocomial infections in
nurseries and hospital wards. S. aureus is a major cause of morbidity and mortality in
haemodialysis patients4 as most patients carry the organism in their anterior nares.
Eradication of nasal carriage of S. aureus may be beneficial in certain clinical conditions
such as recurrent furunculosis. Systemic, in addition to topical, treatment is appropriate
for nasally colonised patients who have infection elsewhere. Topical antibacterial agents
such as mupirocin and chlorhexidine/neomycin are preferred to systemic formulations
when a patient is identified as a carrier.
Nose swabs may be used to investigate carriage of Lancefield group A streptococcus and
Meticillin Resistant Staphylococcus aureus (MRSA)
Specimen Collection, Transport and Storage
Safety considerations
 Use aseptic technique.
 Collect specimens in appropriate CE marked leak proof containers and transport
specimens in sealed plastic bags.
 Collect swabs into Amies transport medium with charcoal22 and transport in
sealed plastic bags.
 Compliance with postal and transport regulations is essential.
 Collect specimens before antimicrobial therapy where possible.
 Specimens should be transported and processed as soon as possible.
 If processing is delayed, refrigeration is preferable to storage at ambient
temperature20. Delays of over 48hr are undesirable.
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Correct specimen type and method of collection
 Plain sterile cotton wool swab. Sample the anterior nares by gently rotating the
swab over the mucosal surface.
 Unless otherwise stated, swabs for bacterial and fungal culture should then be
placed in amies transport medium with charcoal.
 Numbers and frequency of specimen collection are dependent on clinical
condition of patient.
Specimen Processing
Inoculate each agar plate with a swab For the isolation of individual colonies, spread
inoculum with a sterile loop.
Table 11: Culture media, conditions and organisms for all specimens
Staphylococci Identification
Staphylococci are the bacteria most frequently associated with the production of pus.
Staphylococci grow well aerobically on blood agar and form opaque white to cream
colonies, 1–2mm in diameter, after overnight incubation. They are unique in growing on
media with a high salt content, such as MSA. They can be differentiated from
streptococci by their morphology and by the production of catalase. Catalase production
by staphylococci is shown by placing it in a drop of 3% hydrogen peroxide onto the
colonies deposited on a clean glass slide. The appearance of bubbles of oxygen is an
indication of catalase production.
For clinical purposes, staphylococci can be divided into those that produce coagulase and
those that do not. The coagulase-producing staphylococci belong to the species S. aureus,
which is the species of greatest medical interest.
Of the several coagulase-negative species, only two will be considered here—S.
epidermidis and S. saprophyticus.
Although S. aureus is part of the commensal microbial flora of the nose (40% of healthy
adults are positive), skin, and intestinal tract, this species causes impetigo, boils,
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abscesses, wound infection, infection of ulcers and burns, osteomyelitis, mastitis (breast
abscess), pleural empyema, pyomyositis, toxic shock syndrome, and other types of
pyogenic infection. S. epidermidis is also a common commensal of the skin, nose, and
other mucous membranes and possesses a very low pathogenicity. However, its presence
in pus should not always be dismissed as skin contamination. Despite its low infectivity,
S. epidermidis can cause skin infections at the site of an in-dwelling catheter, cannula, or
other device. Infections with S. epidermidis are particularly troublesome in cardiac and
orthopaedic surgery involving the insertion of prosthetic devices (artificial heart valves or
artificial hips).
S. saprophyticus is recognized to be a common cause of urinary tract infections in young
women, being second only to E. coli in some populations.
The distinctive features of the three main species of Staphylococcus are given in Table (
1).
Table 12: Differentiation of medically important species of Staphylococcus
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Figure 25: A flow diagram for the preliminary identification of staphylococci
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In view of the importance of the coagulase test in the identification of S. aureus, this test
is described here in detail. Coagulase is an enzyme that causes plasma to clot.
Staphylococcal coagulase exists in two forms: bound coagulase or clumping factor,
which is demonstrated in the slide test, and free coagulase, which is demonstrated in the
tube test.
• Slide test. On a clean slide, emulsify one or a few similar colonies of staphylococci in a
drop of saline. The suspension must be fairly thick. Dip a straight wire into plasma and
use this to stir the bacterial suspension.
Observe for clumping within 10 seconds. False-negative slide tests occur approximately
10% of the strains of S. aureus. If the slide test is negative for an isolate that seems to be
pathogenic on other grounds (pigment, clinical source), it should be re-examined in the
tube test.
• Tube test. Dispense a few drops (0.5 ml) of plasma into a sterile 12 \ 75 mm tube, and
add two drops of the pure culture in broth. A suspension with an equivalent density may
also be prepared directly from growth on blood agar. Incubate the tube at 35 ∞C for 4–18
hours and then examine for clotting.
The plasma used in the coagulase test may be fresh human or rabbit plasma obtained with
ethylenediamine tetraacetic acid (EDTA). It should be stored in the refrigerator in small
amounts (1 ml), and its performance checked with cultures of S. aureus and S.
epidermidis, run in parallel.
Antimicrobial susceptibility testing
Prudent use of antimicrobials according to local and national protocols is recommended.
Reporting Procedure
Culture
Report presence or absence of specific pathogens, also report results of supplementary
investigations:
Negatives
"Staphylococcus aureus NOT isolated".
"Lancefield group A streptococcus NOT isolated".
Positives
"Staphylococcus aureus isolated".
"Lancefield group A streptococcus isolated".
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Module 4
Session 3: Smart tests, preparation of media and staining smears
Objectives
At the end of this session participants will be able to:
1. Discuss Helicobacter pylori Serum/Whole Blood
2. Explain Salmonella typhi Antigen
3. Explain Vibrio cholerae Rapid visual antigen detection test
4. Explain Preparation of Media
5. Discuss staining smear
Trainers Preparation:




Review the reading material and the session plan.
Prepare the presentation as appropriate and as recommended in the method
column of the session plan, or write the information on a flipchart or board where
all participants can see it.
Prepare copies of the reference materials/handouts and exercises.
Arrange the training room.
Methods and activities

Mini-lecture, question & answers, discussion-lecture
Evaluation/assessment
Questions and answers, trainer’s observation
Estimated Time
195 minutes
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Session plan -3Objectives
4.3.1. Discuss Helicobacter
pylori Serum/Whole Blood
Content





4.3.2. Explain Salmonella
typhi Antigen




4.3.3.Explain Vibrio
cholera Rapid visual
antigen detection test





4.3.4. Explain Preparation
of Media




Methodology
Mini -lecture
Introduction
Principle of the test
30 minutes
Materials and
reagents
Specimen collection
and storage
Procedure
Principle of the test
Question & answers
Materials
Specimen collection 30 minutes
and storage
Procedure
Discussion- lecture
Principle of the test
Materials
45 minutes
Specimen collection
and storage
Procedure
Interpretation and
reporting of results
Blood agar
Chocolate agar
MacConky agar
Mueller-Hinton
agar
Discussion- lecture
60 minutes
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
4.3.5.Discuss staining
smear
Cary-Blair, Amies,
and Stuarts medium
Mini-lecture


Gram’s Stain
Ziehl-Neelsen’s
stain (for acid fast
bacilli)
30 minutes
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6.6 Smart tests
6.6.1 Helicobacter pylori Serum/Whole Blood
The exact role that H. pylori plays in gastrointestinal disease still needs to be precisely
defined and is the subject of ongoing research. However, the prevalence rates for H. pylori
infection as demonstrated by histological and bacteriological methods can approach 90%
in patients who present clinical symptoms of the gastrointestinal diseases listed below
(clinical significant). H. pylori does not appear to invade the bloodstream since no isolates
yet have been detected using commercial blood culture methods. H. pylori infections occur
in human populations throughout the world. In developed countries, about 50% of the
population may have H. pylori infection by the age of 60 years, while only 10-20% of
adults in the third decade of life have it.
In patients who present clinical symptoms relating to the gastrointestinal tract there are two
major methods of investigation: invasive and noninvasive. Invasive methods include
culture of gastric biopsy samples, histological examination of stained biopsy specimens,
or direct detection of the urease activity in the biopsy (CLO test). These methods need to
obtain a biopsy sample by endoscopy, which is expensive, and usually results discomfort
and risk to the patient. Noninvasive techniques include urea breath tests and serological
methods. Urea breath test requires the use of a small amount of radioactivity and a mass
spectrometer. Serologic tests are employed to detect antibodies as human immune response
to H. pylori. Two methods appear to be of great interest regarding their use in H. pylori
routine serology, namely the ELISA and the Western immune blot because they offer the
most versatility in regards to immunoglobulin specificity and relative ease of use.
This H. pylori rapid test detects IgG antibodies specific to H. pylori infection in patient’s
blood or serum. It is a noninvasive method and does not use radioactive isotopes; the
assay procedures are easy and do not require professional training; it provides a rapid
result. It is a useful onsite aid in the diagnosis of H. pylori infection.
CLINICAL SIGNIFICANT
Helicobacter pylori has been associated with a variety of gastrointestinal diseases
including gastritis, duodenal and gastric ulcer, non-ulcer dyspepsia, gastric
adenocarcinoma and lymphoma.
PRINCIPLE OF THE TEST
This assay is a double antigen chromatographic lateral flow immunoassay. The test strip
in the device includes: 1) a burgundy-colored conjugate pad containing colloidal gold
coupled with H. pylori antigens, and 2) nitrocellulose membrane containing a test line (T
line) and a control line (C line). The T line is coated with H. pylori antigens, and the C
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line is coated with goat anti-H. pylori antibody. The antigens used in this device are from
H. pylori cell lysate.
When IgG antibodies specific to H. pylori are present in the specimen, the T line will
become a burgundy-colored band. If antibodies to H. pylori are not present or are present
below the detectable level, no T line will develop. The C line should always appear as a
burgundy-colored band regardless of the presence of antibodies to H. pylori. The C line
serves as an internal qualitative control of the test system to indicate that an adequate
volume of specimen has been applied and the flow occurred.
MATERIALS AND REAGENTS
• Test devices, each sealed in a pouch with a dropper pipette.
• 1 bottle of wash buffer-7 ml PBS diluents with 0.02% sodium azide as a preservative.
• 1 package inserts (Instruction for Use).
• Lancet or other blood collection device (Vacutainer blood collection tubes, sterile
needles and syringes).
• Timer.
Do not expose the kit to temperatures over 30°C (86°F).
SPECIMEN COLLECTION AND STORAGE
1- Serum
• Follow standard laboratory procedures to collect serum specimens.
• Serum specimens can be stored at (9-30) °C for 8 hours, at (2-8) oC for one week, and at
(≤ -20) °C or lower for long term storage. Repeatedly frozen and thawed specimens are
not recommended for this assay.
• Any sediment in serum specimens should be removed by centrifugation. Avoid using
any turbid specimens, which may be contaminated by microorganisms.
2- Whole Blood
• Finger stick sampling is recommended for this assay.
• Middle or ring finger is the preferred puncture site.
• Clean patient’s finger with an alcohol swab. Wait until it is dry.
• Puncture the fingertip with the lancet. Wipe away first sign of blood.
• Gently rub the hand from palm to finger to help form a drop of blood over the punctured
site.
• Use the provided pipette to pick up the blood, and apply one drop of the blood to the
sample well of the device. Then, follow the procedure.
PROCEDURE
1. Refrigerated specimens and other test materials, including devices, must be
equilibrated to room temperature before testing.
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2. Remove the device from its wrap pouch prior to performing the assay, label the device
with identification.
3. Add one drop of fresh blood or serum to the sample well marked "S". Allow about 30
seconds for the specimen to be absorbed totally. Discard the first three drops of wash
buffer from the wash buffer squeeze bottle. Then add three drops of wash buffer into the
sample well.
4. Strong positive results may be observed in 2-3 minutes or depending on manufacturers.
Weak positive results may take a longer time, up to 7 minutes. For the whole blood test, a
slight hemolysis might be observed, but it does not interfere with the results.
INTERPRETATION OF RESULTS
IMPORTANT: Interpret the results depending on manufacturers. The T line should
always be interpreted independently of the C line.
Figure 26: Interpretation of Smart Test Results
Positive:
If both the C line and T line appear, the result indicates that the IgG antibodies specific to
H. pylori are detected and the result is positive.
A faint line in test region indicates a borderline specimen, which should be re-tested
using an alternative method for confirmation.
Negative:
If only the C line appears in the control region, the test indicates that no antibodies to H.
pylori are detected and the result is negative.
Invalid:
When no control line appears within 5 minutes, repeat the test with a new test device.
QUALITY CONTROL PROCEDURE
• Built-in Control Features
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This test contains a built-in quality control feature, the C line. The appearance of the
burgundy C line indicates that that an adequate volume of specimen and wash buffer has
been applied and the flow occurred.
• External Quality Control
External controls are recommended, positive and negative, to monitor the performance of
the assay.
6.6.2 Salmonella typhi Antigen
Immunochromatographic rapid assay for the Detection of Salmonella Typhi Antigen in
Human Stool or Serum Specimens, Typhoid fever is a life threatening illness caused by the
bacterium Salmonella typhi, and was observed by Eberth (1880) in the mesenteric nodes
and spleen of fatal cases of typhoid fever. It is common in developing countries where it
affects about 12.5 million persons annually. The infection is acquired typically by
ingestion. On reaching the gut, the bacilli attach themselves to the epithelial cells of the
intestinal villi and penetrate to the lamina and submucosa. They are then phagocytosed
there by polymorphs and macrophages. The ability to resist intracellular killing and to
multiply within these cells is a measure of their virulence. They enter the mesenteric lymph
nodes, where they multiply and, via the thoracic duct, enter the blood stream. A transient
bacteremia follows, during which the bacilli are seeded in the liver, gall bladder, spleen,
bone marrow, lymph nodes and kidneys, where further multiplication takes place. Towards
the end of the incubation period, there occurs a massive bacteremia from these sites,
heralding the onset of the clinical symptoms. The diagnosis of typhoid consists of isolation
of the bacilli and the demonstration of antibodies. The isolation of the bacilli is very time
consuming and antibody detection is not very specific. Other tests include the Widal
reaction, has developed a test that takes only 10-20 minutes and requires only a small
quantity of stool or one drop of serum to perform. It is the easiest and most specific method
for detecting S. typhi infection.
PRINCIPLE OF THE TEST
Cortez One Step S. Typhi Antigen test is a qualitative one step immunochromatographic
assay. The test employs a conation of monoclonal antibody/colloidal gold dye conjugate
and a polyclonal antibody immobilized on the solid phase. This will selectively identify S.
typhi antigen associated S. typhi (typhoid) infection with a high degree of sensitivity and
specificity.
As the specimen flows through the absorbent pad in the sample well and through the
antibody/colloidal gold complex any S. typhi antigen present in the sample binds to the
conjugate forming an antigen/antibody complex. The sample and dye complex continue to
migrate along the membrane to the immobilized polyclonal antibody. In the presence of S.
typhi, the polyclonal antibody captures the complex. This forms a visible pink/purple band
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in the (B) or test area of the card. If no antigen is present, there is no line formation in the
(B) area. The remaining complex continues to migrate to another immobilized antibody on
the membrane in the (C) or Control area of the card, and is captured which then forms a
band indicating proper performance of the test.
MATERIALS
Each kit contains:
1. Cortez S. typhi Antigen test. Each cassette contains a test strip with S. typhi specific
antibody on the test region of the membrane and colored S. typhi antibody-gold conjugate
pad.
2. Fecal sample buffer – 2 bottles, 8 mL each.
3. Instruction for use.
4. Specimen collection container.
5. Timer.
SPECIMEN COLLECTION AND STORAGE
Cortez One Step S. typhi Antigen test can be run on stool or serum samples. The test works
best on fresh samples. If testing cannot be done immediately, they should be stored at (28)°C after collection for up to 3 days. If testing cannot be done within 3 days, serum can
be stored frozen at (–20) °C or colder. Shipment of samples should comply with local
regulations for transport of etiologic agents.
Stool and serum specimens should be collected in containers that do not contain media,
preservatives, animal serum or detergents as any of these additives may interfere with the
Cortez S. typhi Antigen Test.
REAGENT PREPARATION
Bring all reagents, including test device, to room temperature (20-30) oC before use.
STOOL SPECIMEN PREPARATION
Add about 1/4 gram stool to approximately 500μl of the extraction reagent provided (about
12 drops from the dropper vial provided). Mix well and allow to sit for 5 minutes or so to
allow the large particles to settle.(amount of stool, extraction reagent and appropriate
wetting time depending on manufacturer).
Note: Watery or diarrhea specimens are inappropriate for testing.
PROCEDURE
1. Bring all materials and specimens to room temperature (8 – 30) °C.
2. Remove the test card from the sealed foil pouch.
3. For stool samples: use the provided pipette to transfer sample from the upper layer of
the stool extract and add 3 drops to the sample well (marked as “A”).
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4. For serum samples: use the provided pipette to transfer the serum sample and add 3
drops to the sample well (marked as “A”).
5. Read the result at 20 minutes. A strong positive sample may show test band earlier.
However, to confirm a result is negative, it must wait 20 minutes to read the results.
Note: The amount of S. typhi antigens present in serum is typically less than that in
stool. This may decrease the sensitivity of the test when using serum depending how
soon after the onset of the infection the test is performed. Early infection typically
exhibits greater levels of the antigen in the serum than in later infection. To confirm
serum results:
The use of a stool sample is recommended if serum is used first and a negative
result is obtained and typhoid is still suspected, A second test run on a stool sample
should be performed.
INTERPRETATION OF RESULTS
Positive result: A distinct pink colored band appears on test line regions, in addition to a
pink line on the control line region.
Negative result: No line appears in the test line region. A distinct pink line shows on the
control line Region.
Invalid: The control line next to the test line does not become visible within 20 minutes
after the addition of the sample.
Figure 27: Interpretation of Salmonella Smart Test
QUALITY CONTROL
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1. The control band is an internal reagent and procedural control. It will appear if the test
has been performed correctly and the reagents are reactive.
2. Good Laboratory Practice recommends the daily use of control materials to validate the
reliability of the device.
LIMITATIONS
1. The test is for qualitative detection of S. typhi antigen in stool or serum sample and does
not indicate the quantity of the antigens.
2. The test is for in vitro diagnostic use only.
3. For samples that test positive (reactive) by Cortez S. typhi Antigen Test, more specific
confirmatory testing should be done. A definitive clinical diagnosis should only be made
by the physician after all clinical and laboratory finding have been evaluated. The use of a
rapid test alone is not sufficient to diagnose S. typhi infection even if antigen is present.
Also, a negative result does not preclude the possibility of infection with S. typhi.
Specificity
Negative stool samples from patients in areas where typhoid is relatively rare and would
yield a typical negative population, showed no false positives when the test was read within
20 minutes as specified.
Samples that were positive for s. paratyphi however were also negative as the antibodies
used in the S. typhi rapid test are specific for S. typhi only.
6.6.3 Vibrio cholerae Rapid visual antigen detection test
Vibrio cholerae strains belonging to the O1 and O139 serogroups are capable of causing
epidemic and pandemic cholera. The O1 serogroup is subdivided into two serotypes,
Ogawa and Indaba. Serogroup O139, which appeared in India in 1992, has spread rapidly
throughout Asian countries and is considered to be the potential eighth pandemic strain of
cholera. Prompt diagnosis of cholera is of key importance to initiate effective therapy and
to institute proper epidemiological measures. There are definitive indications that the
incidence of this serogroup is on the rise in India and Bangladesh.
Several rapid diagnostic tests for cholera have been described. Some detect the cholera
toxin. The others detect the lipopolysaccharide (LPS) antigen of V. cholerae O1 or O139.
Recently, a multistep colloidal gold-based colorimetric immunoassay known as SMART
was also developed for direct detection of V. cholerae O1 or V. cholerae O139 in stool
specimens and has demonstrated 95% sensitivity and 100% specificity for O1 strains and
100% sensitivity and 97% specificity for O139 strains.
Clinical Significant
Clinically, cholera may range from asymptomatic colonization to sever diarrhea with
massive fluid loss, leading to dehydration, electrolyte disturbances, and death.
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Principle
This test is an immunochromatographic rapid visual antigen detection test for V. cholerae
from stool specimens. The nitrocellulose membrane is coated with monoclonal antibodies
to V. cholerae O1 and O139 lipopolysaccharide (LPS) as two distinct bands. When the test
sample migrates through the nitrocellulose membrane, colloidal gold particles coupled with
anti-V. cholerae O1 and O139 LPS monoclonal antibodies bind to the respective antigens
from the test sample, if present. This antigen-antibody complex moves through the
nitrocellulose membrane and binds to the corresponding immobilized antibodies against V.
cholerae O1 and/or O139, forming a magenta red color band, which confirms positive
results. A control test band should always appear, irrespective of a positive or negative test
sample, to validate the test procedure.
Materials and equipment
In countries at risk for epidemics of cholera, the laboratory’s first role is to be prepared for
a possible epidemic. It must have ready or have access to the supplies and equipment
necessary to identify V. cholerae O1 and/or O139 even in remote locations. Contact the
local laboratory, medical center, or field site where testing will be performed in advance to
ensure all necessary supplies, equipment, and personal protective equipment (PPE) will be
readily available.
Supplies and equipment necessary to perform the rapid test Included with the rapid test kit:

Individually packaged dipsticks in aluminum pouch with desiccant Plastic,
individual, single-use 5-mL test tubes for sample.
 Disposable transfer pipettes (plastic droppers) for stool.
 Foam test tube stand.
 Sterile normal saline or distilled water Disposable, clean sample containers for stool
specimen collection.
 Plastic rack of appropriate size for test tubes (suggested).
 Watch or timer.
 Biosafety supplies/personal protective equipment (PPE):Gloves Goggles or face
shield
 Lab coat.
 Biohazard receptacle for infectious waste.
 Disinfectant.
Storage
The rapid test kit may be stored at ambient temperature (24–26) oC but it is recommended
that the kit be stored at 4oC when not in use for 6 months or more (depending on
manufacturer).
Specimen’s collection
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Fecal specimens should be collected in the early stages of any enteric illness, when
pathogens are usually present in the stool in highest numbers, and before antibiotic therapy
has started.
Collect stool specimens from persons who are suspected of being part of an outbreak and
who meet the following criteria:
1. Currently have “rice water stool.”
2. Had onset of illness less than 4 days before sampling.
3. Have not received antimicrobial treatment for the diarrheal illness.
Collect stools from patients in clean (no disinfectant or detergent residue) containers with
tight-fitting, leak-proof lids. Do not collect specimens from bedpans, as they may contain
residual disinfectant or other contaminants. Stool should be refrigerated if possible and
processed within a maximum of 2 hours after collection.
Suitable specimens:
Liquid fecal specimens, ideal “rice water stool”—clear, watery stools, likened to water
from boiled rice, an appearance classically seen in cholera cases.
Viscous, mucoid, or semisolid stools—acceptable but must be diluted before use with
normal sterile saline or sterile distilled water.
Unacceptable specimens:
Solid stools—not recommended, as particulate matter from these specimens may clog the
nitrocellulose membrane restricting migration of the sample.
Rectal swabs—the dipstick assay has not been evaluated for use with rectal swabs.
Procedure
1. Put on personal protective equipment (PPE):
lab coat, gloves, goggles and face shield. Wear
PPE at all times while handling the specimen
2. Collect stool sample in plastic cup. Label cup
with patient identifier.
3. Remove 1 test tube from kit and place test
tube in Styrofoam or plastic rack. Label the test
tube.
4. Remove 1 plastic dropper from kit.
5. Fill dropper halfway (150–200 µL) with stool
sample, and transfer to test tube. Do not put
more liquid than this amount in the test tube.
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6. Open packet with rapid test, Label the dipstick
with patient identifier.
7. Place the dipstick in the test tube with the
arrows facing DOWN. The end of the strip
should be submerged in the stool. The arrows
should remain ABOVE the level of the stool.
8. Wait 15–20 minutes for the test to complete.
When complete, you will see a band near the top control band
of the dipstick (internal control band).
9. Review and interpret results within 15–20
minutes: A. Positive, V. cholerae O1 B. Positive,
V. cholerae O139 C. Positive, both V. cholerae
O1 and O139 D. Negative for V. cholerae O1
and O139 E. Test did not work correctly (invalid
test) Do not interpret results after 30 minutes.
A
B
C
D
E
10. Place all waste in a double-lined plastic bag
labeled “biohazard.”
Figure28: Rapid Test Instruction Sheet, for a graphic representation of the test
procedure
1. Allow time for the kit to come to room temperature before opening and using.
2. Label a clean, unused test tube with the patient identification code. Transfer 150– 200
L of the patient’s liquid stool specimen to this test tube.
Note: Viscous, mucoid, or semisolid specimens should be diluted with sterile saline or
distilled water before transfer to the test tube.
3. Tear open aluminum pouch at notched area. Remove the dipstick.
4. Label the dipstick with the patient identification code; take care to not touch the area
marked with an arrow (dipping area).
5. Carefully place the dipstick in the test tube with stool with the arrows facing DOWN.
The end of the strip should be submerged in the stool. Be sure the arrows remain ABOVE
the level of the stool.
6. Leave the dipstick undisturbed for 10–15 minutes or depending on manufacturers.
7. Within 15-20 minutes, a magenta red internal “control” band will appear indicating a
valid test; this line indicates that the reaction is complete. The presence of at least one or
two additional magenta red lines signifies the sample is positive for V. cholerae O1
and/or V. cholerae O139.
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8. Read and record the result of the test within 15–20 minutes. Do not interpret results
after 30 minutes. (Reading & interpretation the result depending on manufacturers).
Interpretation and reporting of results
Interpret test results of each dipstick according to the manufacturer’s instructions.
Guidelines for interpreting test results can be found in Table (1-1).
Record all patient test results in a worksheet or notebook as a permanent record. Record
results as positive, negative, or invalid. If a test needs to be repeated because of an invalid
test result, record the first result (invalid), resolve the problem, and record the repeated
result. Report positive and negative test results to the appropriate person in a timely
manner. The report should include any comments or information the person needs to
properly evaluate the test results.
Table 12: Expected Test Result Outcomes for the Dipstick and Interpretation of
Test Results
Dipstick Test Result
Interpretation
POSITIVE (Figures 3-1 and 3-2,
column A): Appearance of two bands,
one for the control test band and one
test band specific for V. cholerae O1
(Vc O1).
Indicative that the sample is positive only for V.
cholerae O1.
POSITIVE (Figures 3-1 and 3-2,
column B): Appearance of two bands,
one for the control test band and one
test band specific for V. cholerae O139
(Vc O139).
Indicative that the sample is positive only for V.
cholerae O139.
POSITIVE (Figures 3-1 and 3-2,
column C): Appearance of three bands,
one for the control test band, one test
band specific for V. cholerae O1 (Vc
O1) and one test band specific for V.
cholerae O139 (Vc O139)
Indicative that the sample is positive for both V.
cholerae O1 and O139.
NEGATIVE (Figures 3-1 and 3-2,
column D): Appearance of only the
control test band.
Indicative that the sample is negative for V.
cholerae O1 and O139.
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INVALID (Figures 3-1 and 3-2,
column E): Appearance of no control
test band
Indicates a procedural error, deterioration of the
specimen/dipstick or the presence of particulate
matter in the specimen preventing flow of fluid
through the nitrocellulose membrane.
Quality control
Common source of error
Specimens that are not classified as “watery” may clog the nitrocellulose membrane strip,
limit migration of the liquid, and prevent the dipstick reaction resulting in an invalid test.
These specimens should be either diluted with sterile saline or distilled water or rejected as
unacceptable specimens for testing.
A
B
C
D
E
Figure 29:Visual Interpretation of Rapid Dipstick Test Results
Precautions
Be sure to run the test exactly as instructed, adhering to all procedures. Use sterile
technique throughout the test procedure. Wear gloves, goggles and face shield, and a lab
coat when handling and testing all clinical specimens.
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6.7 Preparation of Media
With just a few diagnostic materials, a laboratory can make an important contribution to
individual patient care through accurate etiological diagnosis.
In most developing countries bacteriological laboratory practice is hampered by a
shortage of culture media and basic reagents, which are very costly to import. However,
the number of culture media and reagents that have to be purchased can be reduced to the
essential ones, through rational selection, as has been the case with essential drug lists.
Additionally, some simple media
and reagents can be produced or prepared locally.
Each batch of medium prepared from individual ingredients or each different
manufacturer’s lot number of commercial dehydrated medium and each batch of medium
prepared from individual ingredients should be quality controlled before use.
Immediately after preparation, each medium should be tested, as appropriate, with a
reference strain .
for proper growth characteristics as described for each medium.
A record of all media preparation or purchase dates and quality control test results should
be kept, and any unusual characteristic (e.g., the color of the medium or slow growth of
test bacteria) should be noted. Many media call for the use of defibrinated blood.
Agar media should be dispensed into 15 x 100-mm or 15 x 150-mm Petri dishes to a
uniform depth of 3–4 mm; approximately 20-ml of liquid agar medium will achieve this
depth in a 15 x 100-mm plate. If agar is cooled to 50°C prior to pouring, condensation is
minimized. After pouring, the plates should be kept at room temperature for several hours
to prevent excess condensation from forming on the covers of the dishes. Another means
by which condensation will be reduced is if plates are stacked so that they cool more
slowly. Alternatively, if when
preparing selective media (e.g., MacConkey [MAC], xylose lysine desoxycholate [XLD],
thiosulfate citrate bile salts [TCBS] agar, etc.), conditions are such that there is little
chance that the cooling media will be contaminated, after the agar is poured into the
plates, the lids can be placed on the dish so that a small opening is left to let the heat out,
resulting in the formation of less condensation on the upper lid; the lid should remain
slightly open like this for approximately 30 minutes,
while the agar solidifies. If, however, it is likely that the agar will be contaminated if the
lid is left partly open, the agar should be allowed to solidify with the lid closed.
Note: Covering the agar while it is still hot will allow for the formation of a substantial
amount of condensation on the upper lid. If the plates contain condensation, the plates
should be covered at room temperature for 24 hours to allow the condensation to
evaporate. After condensation has evaporated, the plates should be placed in an inverted
position and stored in a plastic bag in an inverted position at 4°C.
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6.7.1 Blood agar: TSA with 5% sheep blood
Sheep blood agar is used as a general blood agar medium, and consists of TSA plus 5%
sheep’s blood. The sheep blood agar plate should appear a bright red color. If the plates
appear dark red, the blood has been added when the agar was too hot; if this happens, the
medium should be discarded and a new batch prepared.
A. Prepare TSA according to the instructions given on the label of the dehydrated
powder. For convenience, 500 ml of molten agar can be prepared in a l-liter flask.
Add 20 g of agar into 500 ml of water. Heat to dissolve.
B. Autoclave at 121°C for 20 minutes. Cool to 60°C.
C. Add 5% sterile, defibrinated sheep blood (i.e., add 25 ml sheep blood to 500 ml of
agar). If a different volume of basal medium is prepared, the amount of blood
added must be adjusted accordingly to 5% (e.g., 50 ml of blood per liter of
medium).
D. Dispense 20 ml into 15 x 100-mm Petri dishes. Allow the medium to solidify and
dry out, place in a plastic bag, and store at 4°C.
Quality control: Test each new, freshly prepared or purchased batch of blood agar plates
for growth and hemolytic reaction with a strain of S. pneumoniae. The colonies are small
and should appear grey to grey-green surrounded by a distinct greenish halo in the agar.
6.7.2 Chocolate agar with TSA base and growth supplement
Chocolate agar with growth supplements is a medium that supports the special growth
requirements needed for the isolation of fastidious organisms (when incubated in a 5%
CO2 atmosphere). Chocolate agar contains a reduced concentration of agar, which
increases the moisture content of the medium.
Supplemented chocolate agar should support the growth of H. influenzae.
Chocolate agar slants for transport and short-term storage can be prepared in the same
manner as that described for agar plates, except that the medium is dispensed in 16 x 125mm screw-cap tubes and slanted before solidifying.
A. Use TSA as the basal medium. Prepare double strength (i.e., 20 g in 250 ml
distilled water). Autoclave and then cool to 50°C. Use the thermometer to verify
the cooling temperature.
B. Prepare a solution of 2% hemoglobin (i.e., 5 g in 250 ml distilled water). Mix the
hemoglobin in 5–6 ml of the distilled water to form a smooth paste. Continue
mixing as the rest of the water is added. Autoclave, and cool to 50°C.
C. Add the hemoglobin solution to the double-strength TSA and continue to hold at
50°C.
D. • Alternative to steps a-c: If a hemoglobin solution is unavailable, an alternative is
to add 5% sterile defibrinated sheep, rabbit, guinea pig, or horse blood (i.e., 5 ml
blood per 100-ml agar) to full-strength TSA (i.e., 20 g in 500 ml distilled water).
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DO NOT use human blood. After the base medium has been autoclaved and
cooled to 50°C, add the blood and place in
E. a hot water bath at no more than 80°C for 15 minutes or until a chocolate color is
achieved. Then cool to 50°C.
F. After the hemoglobin solution or the defibrinated blood has been added to the
base medium and the medium has cooled to 50°C, add growth supplement (e.g.,
IsoVitaleX or Vitox) to a final concentration of 1%.Mix the ingredients by gently
swirling the flask; avoid forming bubbles. Dispense 15–20 ml in each 15 x 100mm Petri dish.
Quality control: All freshly prepared or purchased chocolate agar media should be tested
to determine the medium’s capacity to support growth of the bacteria to beisolated,
particularly H. influenzae. If the medium does not support the growth ofone or all of the
bacteria, the medium should be discarded, and a new batch of medium should be
prepared or purchased.
• Chocolate agar should look brown to brownish-red in color. N. meningitidis
and H. influenzae should appear as a greyish, almost translucent film on the slant’s
surface with no discoloring of the medium after 24 hours of incubation; S. pneumoniae
should appear as small grey to grey-green colonies with a very distinct greenish
discoloring of the medium .
If H. influenzae does not grow, the growth supplement (IsoVitaleX or its equivalent)
may have been inadvertently omitted.C
6.7.3 MacConkey agar (MAC)
MacConkey agar (MAC) is a differential plating medium recommended for use in the
isolation and differentiation of lactose-nonfermenting, gram-negative enteric bacteria
from lactose- ermenting organisms. Colonies of Shigella on MAC appear as convex,
colorless colonies about 2–3 mm in diameter. S. dysenteriae 1 colonies may be smaller. S.
Typhi colonies are flat, colorless and usually 2–3 mm in diameter. Several commercial
brands of MAC are available.Most manufacturers prepare several formulations of MAC,
which may vary in selectivity and thereby affect the isolation of Shigella. For example,
some formulations of MAC do not contain
crystal violet, a selective agent; these types are not as selective and should not be used for
isolation of Shigella. Oxoid MacConkey Agar No. 3, Difco Bacto MacConkey Agar, and
BBL MacConkey Agar are all suitable.
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A. Prepare MAC according to manufacturer’s instructions. [Note: MAC can also be
prepared from individual ingredients, but this produces more lot-to-lot variation
than preparation of a commercially available dehydrated formulation.]
B. Sterilize the medium by autoclaving at 121°C for 15 minutes.
C. Cool to 50°C and pour into Petri plates (to a uniform depth of 3–4 mm).
D. Leave lids ajar for about 20 minutes so that the surface of the agar will dry. Close
lids and store at 4°C for up to 1 month. If plates are to be stored for more than a
few days, put them in a sealed plastic bag to prevent drying.
Quality control: For quality control of MAC, the following organisms should be
adequate for confirmation of selective and inhibitory growth characteristics:
• E. coli should produce pink to red colonies with good to excellent growth; and,
• S. flexneri should produce colorless colonies with fair to good growth, but S.
dysenteriae 1 colonies may be smaller.
6.7.4 Mueller-Hinton agar
Mueller-Hinton agar is the NCCLS-recommended medium used for standardized
antimicrobial susceptibility testing of certain bacteria; the organisms in this document for
which it is appropriate to use this formulation of Mueller-Hinton medium (i.e.,
unsupplemented Mueller-Hinton) are S. Typhi, Shigella spp., and V. cholerae.
[Note: Several formulations of Mueller-Hinton agar are commercially available.
This laboratory manual suggests that Mueller-Hinton agar medium should not be
prepared from individual ingredients because this can diminish the quality. Commercial
dehydrated Mueller-Hinton is carefully quality controlled before
being released for sale.]
A. Follow manufacturer’s instructions to prepare medium.
B. After autoclaving, cool medium to 50°C in a water bath.
C. Measure 60–70 ml of medium per plate into 15 x 150-mm plates, or measure 25–
30 ml per plate into 15 x 100-mm plates. Agar should be poured into flatbottom
glass or plastic Petri dishes on a level pouring surface to a uniform depth of 3–4
mm. Using more or less agar will affect the susceptibility results. Agar deeper
than 4 mm may cause false-resistance results, whereas agar less than 4 mm deep
may be associated with a false-susceptibility report.
D. Freshly prepared plates may be used the same day or stored in a refrigerator (at
2°–8°C) for up to 2 weeks. If plates are not used within 7 days of preparation,
they should be wrapped in plastic to minimize evaporation. Just before use, if
excess moisture is on the surface, plates should be placed in an incubator (35°–
37°C) until the moisture evaporates (usually 10–30 min).
Do not leave lids ajar because the medium is easily contaminated.
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Quality control: Each new lot of Mueller-Hinton agar should be quality controlled before
use by testing the E. coli ATCC 25922 standard strain for antimicrobial susceptibility
testing. (This formulation of Mueller-Hinton agar can also be used for testing of grampositive aerobes, in which case S. aureus ATCC 25923 can be used as a quality control
strain.) The pH of each new lot of Mueller-Hinton should be between 7.2 and 7.4; if
the pH is outside this range, the pH of the medium should not be adjusted by the
addition of acid or base, i.e., the batch of Mueller-Hinton plates should be discarded
and a new batch of plates prepared. If the pH for every batch is too high or low, the
entire lot of dehydrated medium may have to be returned to the manufacturer as
unsatisfactory. Inhibition zone sizes / minimal inhibitory concentration (MIC) values for
quality control are included in the antimicrobial susceptibility testing section of each
pathogenspecific chapter.
6.7.5 Cary-Blair medium, Amies medium, and Stuarts medium
Prepare each of these transport media according to the manufacturer’s instructions.
[Note: Several dehydrated formulations of Cary-Blair are commercially available; some
require the addition of calcium chloride and some do not.] These media can also be
prepared from individual ingredients; however, it is very difficult to make a well qualitycontrolled batch and so this manual recommends purchasing them from a manufacturer.
When the Cary-Blair medium is prepared, it should be dispensed into containers in
sufficient volume so that swabs will be covered by at least 4 cm of medium. For example,
5- to 6-ml amounts may be dispensed into 13 x 100-mm screw cap tubes. With the caps
loosened, sterilize the medium by steaming (not by autoclave) at 100°C for 15 minutes.
Tighten the caps after sterilization, and store the medium at 15°–30°C.
These media are quite stable if stored in tightly sealed containers in a cool dark place so
that the medium does not dry out. Each may be used for up to 1 year as long as no loss of
volume, visible contamination (e.g., foreign objects or bacterial growth), or color change
is observed. Prepared Amies medium that has been stored for longer than 9 months,
however, should be freshly steamed and the charcoal re-suspended before use.
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6.8 Staining Smears
6.8.1 Gram’s Stain
Organisms are classified according to their Gram staining reaction. Gram positive
bacteria have thicker and denser peptidoglycan layers in their cell walls. Iodine penetrates
the cell wall in these bacteria and alters the blue dye to inhibit its diffusion through the
cell wall during decolourisation. Gram positive bacteria must have an intact cell wall to
produce a positive reaction. Gram negative cells which do not retain the methyl/crystal
violet are stained by a counterstain26. Neutral red, safranin or carbol fuchsin may be used
as the counterstain.
Gram ’s Method for examination of smears
1. Prepare a smear and heat gently to fix.
2. Flood the slide with 0.5% methyl/crystal violet and leave for 30 s.
3. Tilt the slide, pour on sufficient (1%) Lugol’s iodine to wash away the stain,
cover with fresh iodine and allow to act for 30 s.
4. Tilt the slide and wash off the iodine with 95 - 100% ethanol or acetone until
colour ceases to run out of the smear.
5. Rinse with water.
6. Pour on 0.1% counterstain (neutral red, safranin or carbol fuchsin) and leave to
act for about 2 min.
7. Wash with water and blot dry.
Interpretation
Positive Result
Gram positive organisms stain deep blue/purple.
Negative Result
Gram negative organisms stain pink/red.
Quality Control Organisms
A culture containing Gram positive and Gram negative organisms may be used for
quality control.
6.8.2 Ziehl-Neelsen’s stain (for acid fast bacilli)
This staining technique is used to demonstrate the presence of acid and alcohol fast
bacilli (AAFB) which have waxy envelopes that make them difficult to stain and
decolourise. Auramine-phenol staining is more sensitive than Ziehl-Neelsen’s and is thus
more suitable for assessment of smears from clinical specimens. Ziehl-Neelsen’s staining
provides morphological details and is more useful for confirming the presence of AAFB
in positive cultures.
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Method
1. Flood the slide with strong carbol fuschin.
2. Heat gently, and once slide is just ‘‘steaming’’ leave for 3-5 min.
3. Rinse well with water.
4. Decolourise for 2-3 min with a (3% v/v) acid-alcohol solution, rinse with water,
then replace with fresh acid-alcohol for 3-4 min until the slide remains a faint
pink colour.
5. Rinse well with water.
6. Counter stain with (1% w/v) methylene blue or malachite green for 30 s.
7. Rinse with water and allow to dry.
8. Apply immersion oil and read with a transmitted light microscope.
Note: Follow manufacturer’s procedure if commercial kits are used.
Interpretation
Positive Result
Acid fast bacilli vary from 0.5-10 μm in length16 and stain red. Some may appear
beaded.
Negative Result
All other organisms and background material stain green if malachite green counterstain
is used or blue if methylene blue counterstain is used.
Quality control
Positive Control
Mycobacterium species.
Negative Control
A proven negative smear may be used as the negative control.
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Module five: Parasitology and biochemistry
Module Objectives:
At the end of this module the participant will be able to:
1. Diagnose malaria parasite
2. Apply biochemistry tests
Modules Sessions


Session 1: Parasitology
Session 2: biochemistry
Evaluation/ Assessment
Questions and answers, participants’ summaries, trainer’s evaluation
Estimated Training Time
5.30 hours
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Module 5
Session 1: Parasitology
Objectives
At the end of this session participants will be able to:
1. Apply method for malaria parasite diagnosis
2. Apply staining blood films with Giemsa stain
Trainers Preparation:




Review the reading material and the session plan.
Prepare the presentation as appropriate and as recommended in the method
column of the session plan, or write the information on a flipchart or board where
all participants can see it.
Prepare copies of the reference materials/handouts and exercises.
Arrange the training room.
Methods and activities

Mini-lecture
Evaluation/assessment
Questions and answers, trainer’s observation
Estimated Time
50 minutes
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Session plan -1-
Objectives
5.1.1 Apply method for
malaria parasite diagnosis.
5.1.2.Apply staining blood
films with Giemsa stain
Content



Method
- Thin film
- Thick film
Methodology
Mini-lecture
25 minutes
Mini-lecture
First method
Second method
25 minutes
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7. Parasitology
7.1 Malaria parasite diagnosis
Malaria is a serious, sometimes fatal disease resulting from infection with Plasmodium
spp. transmitted by the bite of Anopheline mosquitoes.. The clinical diagnosis, where
malaria is suspected based on the history, symptoms and clinical findings must always be
confirmed by a laboratory diagnosis.
Laboratory diagnosis of malaria involves identification of malaria parasite or its
antigens/products in the blood of the patient.
Clinical significance
Malaria parasites are usually detected in blood films stained with Field or Geimsa stains.
They may also be detected using an immunological procedure known as a dipstick test
It is important for the prognosis and treatment of the disease that the species involved are
identified in the laboratory.
Preparation of a thick and a thin blood film on the same slide
There are four species of the genus plasmodium responsible for the malarial parasite
infections that commonly infect man, P.falciparum, P.vivax, P.malariae and P.ovale. The
most important of these is P.falciparum because it can be rapidly fatal and is responsible
for the majority of malaria related deaths
Clinical significance
For routine malaria microscopy, a thin and a thick film are made on the same slide. The
thick film is used for the detection of parasites, while the thin film is used in identifying
the species of parasite
Principal
Blood to be examined for malaria parasites is usually collected at a health centre.
The most suitable time for collection is at the height of an episode of fever, when the
parasites are most numerous in the blood. Blood specimens should always be collected
before anti- malarial drugs are given Cleaning the finger before collecting a capillary
bloodsample Using a lancet to puncture the ball of the finger
METHOD
1. With the patient’s left hand palm upwards, select the third or fourth finger. (The big toe
can be used with infants. The thumb should never be used for adults or children.) Use
cotton wool lightly soaked in ethanol to clean the finger — using firm strokes to remove
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dirt and grease from the ball of the finger, Dry the finger with a clean piece of cotton
wool (or lint).
2. With a sterile lancet, puncture the ball of the finger, using a quick rolling action. By
applying gentle pressure to the finger, express the first drop of blood and wipe it away
with dry cotton wool. Make sure that no strands of cotton wool remain on the finger
3. Working quickly and handling clean slides only by the edges, collect the blood as
follows:
 Apply gentle pressure to the finger and collect a single small drop of blood, about
this size _, on to the middle of the slide. This is for the thin film.
 Apply further pressure to express more blood and collect two or three larger
drops, about this size _, on to the slide about 1 cm from the drop intended for the
thin film Wipe the remaining blood away with cotton wool.
Thin film: Using another clean slide as a “spreader”, and with the slide with the blood
drops resting on a flat, firm surface, touch the small drop with the spreader and allow the
blood to run along its edge. Firmly push the spreader along the slide, away from the
largest drops, keeping the spreader at an angle of 45° Make sure that the spreader is in
even contact with the surface of the slide all the time the blood is being spread.
Thick film: Always handle slides by the edges, or by a corner, to make the thick film as
follows: Using the corner of the spreader, quickly join the larger drops of blood and
spread them to make an even thick film, allow the thick film to dry in a flat, level position
protected from flies, dust and extreme heat. Label the dry film with a grease pencil by
writing across the thicker portion of the thin film the patient’s name or number and date.
Materials and reagents














Microscope
Clean glass microscope slides
Sterile blood lancets
Cotton wool
Grease pencil
Methanol
70% Ethanol.
Measuring cylinders, 10, 50 and 100ml
Beakers, 50 and 250ml
Staining troughs
Glass rods
Wash bottle
Slide forceps
Slide racks
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 Timer
 Giemsa stain
 Methanol in a drop bottle
 Buffered water, pH 7.2 or distilled water
Staining blood films with Giemsa stain
Principle
During staining of the blood film, the haemoglobin in the erythrocytes dissolves(de
haemoglobin ization) and is removed by the water in the staining solution. All that remain
are the parasites and the leukocytes, which can be seen under the microscope.
First method: Rapid method for staining thick and thin blood films
This method is suitable for rapid staining of thick films when urgent results are required.
It uses much more stain than the regular method
1. Using forceps, place the slides back to back in a staining trough
2- Prepare a 3% Giemsa solution in buffered or distilled water, pH 7.2, in sufficient
quantity to fill the number of staining troughs being used, mix the stain well.
3- Pour the stain gently into the staining trough, until all the slides are totally covered.
Stain for 30–45 minutes out of sunlight.
4. Pour clean water gently into the trough to remove the deposit on the surface of the
staining solution
5. Gently pour off the remaining stain), and rinse again in clean water for a few seconds.
Pour the water off.
Second method
1. Allow the thick film to dry thoroughly; if results are required urgently, drying may be
hastened by fanning, or briefly exposing the slide to gentle heat such as that from a
microscope lamp. Care should be taken to avoid overheating, otherwise the thick film
will be heat-fixed
2. Fix the thin film by adding three drops of methanol, or by dipping it into a container of
ethanol for a few seconds. To permit dehaemoglobinization, the thick film should not be
fixed; therefore avoid exposure of the thick film to methanol or methanol vapour
3. Prepare a 10% Giemsa solution in buffered or distilled water, pH 7.2; if a small
quantity is being used, three drops of stain per ml of buffered water will give the correct
concentration of Giemsa solution. One slide requires about 3ml of madeup stain. Mix the
stain well with a glass rod.
4. Gently pour the stain on to the slides or use a pipette. Stain for 5–10 minutes.
5. Gently flush the stain off the slides by adding drops of clean water. Do not tip off the
stain and then wash, as this will leave a deposit of scum over the smears.
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6. Place the slides in the slide rack to drain and dry, film side downwards; making sure
that the film does not touch the slide rack.
Microscopic examination
Examine the slide under the microscope using the ¥100 objective. Malaria parasites found
in the blood are at different stages of development Some malaria parasites have granules
of pigments in their cytoplasm
Thin blood films: In thin blood films, the infected erythrocytes may remain unchanged
or have a different colors or shape, or may contain pink (“Schüffner’s”) or red (“James”)
thin films can be used to identify the species of malaria parasite
Note: In patients who have been suffering from malaria for a long time, monocytes may
be seen in the thin blood film; the cytoplasm often contains brown or greenish black
bodies (siderophils). In patients who have recently received an injection of an
antimalarial drug, the parasites stain poorly and appear distorted and indistinct.
Thick blood films: In thick blood films, the background should be clean and free from
debris, as the infected erythrocytes are lysed. The malaria parasites should have deep red
chromatin and blue or pale purplish-blue cytoplasm. In thick films stained with Giemsa,
the nuclei of leukocytes should be stained dark purple. Schüffner’s dots may be seen
around the malaria parasites. Thick blood films are used for estimating the parasite
density, as described below.
Reporting results
If the result of the examination of the stained blood films is positive, specify:
 the species of parasite found
 the stage of development of the parasite
 the parasite density.
Blood films containing P. ovale and P. vivax may contain few parasites and therefore
take more time to examine under the microscope. However, it is necessary to differentiate
the two species, since they may reappear in the blood without reinfection.
Patients infected with P. ovale or P. vivax require additional treatment to eradicate the
liver stages of these parasites.
A patient can harbour more than one species of malaria parasite at the same time(e.g. P.
falciparum and P. malariae or P. falciparum and P. vivax). If the result is negative, report
as “no parasites found”.
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Figure 30: P. Falciparum
1: Normal red cell , 2-18:Trophozoites (among these) , 2-10:correspond to ring stage
trophozoites , 19-26:Schizonts (26 is ruptured schizont), 27&28: Mature
macrogametocytes , 29&30:Mature microgametocytes (male)
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Figure 31: P. Ovale
1: Normal red cell , 2-5: Young trophozoites (Rings) , 6-15:Trophozoites , 1623:Schizonts , 24:Macrogametocytes( female) , 25: Microgametocytes (male)
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Figure 32: P. Vivax
1: Normal red cell , 2-6: Young trophozoites (Ring stage parasites) , 718:Trophozoites , 19-27:Schizonts , 28&29:Macrogametocytes( female) , 30:
Microgametocytes (male)
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Module 5
Session 2: Biochemistry
Objectives
At the end of this session participants will be able to:
1. Estimate blood glucose level
2. Estimate ALT
3. Estimate AST
4. Estimate Alkaline Phosphatase
5. Estimate total serum bilirubin
6. Estimate renal function test (blood urea, serum creatinine, uric acid)
7. Estimate total cholesterol and triglycerides
8. Estimate HDL Cholesterol LDL Cholesterol
Trainers Preparation:




Review the reading material and the session plan.
Prepare the presentation as appropriate and as recommended in the method
column of the session plan, or write the information on a flipchart or board where
all participants can see it.
Prepare copies of the reference materials/handouts and exercises.
Arrange the training room.
Methods and activities

Mini-lecture, question & answers, discussion, discussion-lecture
Evaluation/assessment
Questions and answers, trainer’s observation
Estimated Time
280 minutes
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Session plan -2-
Objectives
5.2.1.Estimate blood
glucose level
Content





5.2.2.Estimate ALT






5.2.3. Estimate AST






5.2.4. Estimate Alkaline
Phosphatase




Causes of
Hyperglycemia
Causes of
hypoglycemia
Reagents and
materials
Procedure
Reference intervals
Causes of high ALT
Principle
Specimen
Reagents and
materials
Reference value
Limitation
Causes of high AST
Principle
Specimen
Reagents and
materials
Reference value
Limitation
Principle
Specimen
Reagents and
materials
Procedure
Methodology
Question &answers
30 minutes
Mini-lecture
30 minutes
Question & answer
20 minutes
Question & answer
20 minutes
236 | P a g e
5.2.5. Estimate total serum
bilirubin
5.2.6. Estimate renal
function test (blood urea,
serum creatinine, uric acid)
5.2.7. Estimate total
cholesterol and
triglycerides
5.2.8. Estimate HDL
Cholesterol and LDL
Cholesterol

Reference value



Discussion



Principle
Specimen
Reagents and
materials
Procedure
Reference value
Limitation



Blood urea
Serum creatinine
Uric acid
Discussion- lecture

Lipid profile
- Total cholesterol
- Triglycerides

Lipid profile
- HDL Cholesterol
- LDL Cholesterol
30 minutes
60 minutes
Question & answers
45 minutes
Question & answers
45 minutes
237 | P a g e
238 | P a g e
8. Biochemistry
8.1 Blood glucose
Glucose is the chief source of energy in the body. The levels of this compound are
balanced by digestion and absorption of carbohydrates in the intestine, its storage and
release in the liver and its utilisation in the muscle.
Causes of Hyperglycemia:
 Insulin – dependent type (IDDM)
 Non – insulin dependent type (NIDDM)
 non obese NIDDM
 obese NIDDM
 Gestational diabetes
 Impaired glucose tolerance: Glucose intolerance associated with certain
conditions and syndrome such as (chronic pancreatitis with endocrine and
exocrine insufficiency, cushing syndrome , acromegaly )
Causes of hypoglycemia:
 Deficient glucose production
 Ketotic hypoglycemia
 Glycogen storage disorder
 Deficiency of gluconeogenic enzyme
 Galactosemia
 Hereditary fructose intolerance
 Leucine hypersensitivity
 Endogenous hyperinsulinism
Glucose is determined after enzymatic oxidation in the presence of glucose oxidase. The
hydrogen peroxide formed reacts, under catalysis of peroxidase, with phenol and 4aminophenazone to form a red-violet quinoneimine dye. The intensity of the colour
produced is directly proportional to the concentration of glucose in the sample.
Glucose + O2+H2O --------------GOD
H2O2 + Gluconate
2H2O2+ Phenol+ 4-Amino-antipyrine
H2O+ Quinonimine
SPECIMEN
Serum, heparin plasma, or fluoride plasma may be used. Plasma or serum samples
without preservatives should be separated from the cells or clot within a half hour of
being drawn. Glucose in separated, un-hemolyzed serum is stable up to four hours at
25°C and up to 24 hours at 4°C.
REAGENTS AND MATERIALS
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1- Full automated auto analyzer or Spectrophotometer
2- Centrifuge
3- Gel tube (clot activator)
4- Spectrophotometer cuvettes
5- Deionized water
6- Pipettes
7- Glucose standards 20 mg/dL 100 mg/dL, 200 mg/dL, 400 mg/dL
8- Heating block or water bath 37°C
9- Timer
10- Reagents :
Reagent 1
Buffer solution
Reagent 2
tris buffer pH 7
Phenol
glucose oxidase
Peroxidase
4-amino-antipyrine
Reagent 3
standard glucose
Preparation: Dissolve the powder R2 with the Buffer R1
100 mmol/l
0.3 mmol
10 000 u/l
1 000 u/l
2.6 mmol/l
100 mg/dl
PROCEDURE


Wave length ………… 505 nm ( 490 – 550)
Temperature………….37 ċ
Blank
Standard
Sample
Standard
10 µl
Sample
10µl
Working reagent
1 ml
1ml
1ml
 Mix incubate 10 minutes at 37 ċ or 30 minutes at room temperature ( 20 ċ – 30 ċ )
 The color is stable 30 minutes
 Linearity : up to 500 mg / dl
Calculation:
Glucose concentration =O. D. sample / O.D. standard * concentration of standard .
QUALITY CONTROL
Level one and level two serum controls are tested with each patient run. The level one
control range is 70-85 mg/dL and the level two range is 271-306 mg/dL.
RESULTS
Using graph paper, plot the Absorbance on the vertical (y axis) against the concentration
on the horizontal (x axis) for each of the glucose standards.
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1- Draw a "best fit line" and use this standard curve to determine the glucose
concentration for the controls and patient specimens.
2- Verify that the control results are acceptable before reporting patient results.
3REFERENCE INTERVALS
The reference range for glucose is as follows:
Cord
45-96 mg/dL
Premature 20-60 mg/dL
Newborn
40-60 mg/dL
1 wk
50-80 mg/dL
Child
60-100 mg/dL
Adult
74-100 mg/dL
>60 yr
82-115 mg/dL
>90 yr
75-121 mg/dL
LIMITATIONS
1- Serum and plasma must be separated from the red blood cells promptly to prevent
glycolysis. Glucose will decrease approximately 7% per hour when left in contact
with red cells.
2- Whole blood glucose is 12-15% less than serum glucose.
3- Venous blood glucose is approximately 5 mg/dL less than arterial or capillary
blood glucose.
8.2 Liver Function Test
8.2.1 Alanine Aminotransferase (ALT)
Is found mainly in the liver, but also in smaller amounts in the kidneys , heart , muscles,
and pancreas.
ALT is measured to see if the liver is damaged or diseased. Low levels of ALT are
normally found in the blood. But when the liver is damaged or diseased, it releases ALT
into the bloodstream, which makes ALT levels go up. Most increases in ALT levels are
caused by liver damage.
The ALT test is often done along with other tests that check for liver damage, including
aspartate aminotransferase (AST), alkaline phosphatase, lactate dehydrogenase (LDH),
and bilirubin. Both ALT and AST levels are reliable tests for liver damage.
Causes of high ALT:
 Alcohol abuse
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 Medications
 Acute and Chronic hepatitis
Principle:
Kinetic determination of ALT activity , method recommended by IFCC
2-oxoglutarate +l-alanine-----ALT-------- glutamate + pyruvate
Pyruvate + NADH + H+-----------LDH------- l-lactate + NAD+
The rate of NADH consumption is determined photometrically and is direct proportional
to the ALT activity in the sample
SPECIMEN
A. Recommended specimen: 100 μl of serum or heparinised plasma or EDTA
plasma. Collect specimens by standard venipuncture technique. Heparin may be
used as an anticoagulant for plasma specimens. Handle specimens in stoppered
containers to avoid contamination and evaporation. Follow universal precautions
when performing phlebotomy or handling patient specimens, calibrators, or other
serum-based products. Discard contaminated materials with infectious waste.
B. Remove serum promptly from the clot. Haemolysed specimens should not be used
because ALT contamination from red cells will occur. DO NOT freeze the
sample; this will cause a loss of ALT activity.
C. If concentration is greater than the analyser range, dilute with an equal volume of
isotonic saline and re-analyses. Multiply the result by 2 to obtain the original ALT
activity.
REAGENTS AND MATERIALS
A. full automated autoanalyzer or spectrophotometer
B. Centrifuge
C. Gel tube (clot activator)
D. Reagents :
Reagent 1
tris buffer P H 7.5
Buffer reagent
L-alalnine
Reagent 2
NADH
Substrate
LDH
Oxoglutarate
100mmol/l
500 mmol/l
0.18 mmol/l
1200u/l
15 mmol/l
Preparation:
Dissolve one vial of reagent 2 with 3 ml of R 1 buffer, this working reagent stable 13
days at 2 – 8 ċ or 24 hours at 20 – 25 days.
Procedure:
 Wave length ----------340 nm
 Working reagent 1 ml mixed with sample 100 µl and wait 1 minute
242 | P a g e
 Measure the extinction decrease per minute for 1 – 3 minutes
Calculation :
Δ OD/min * 1750 = u/l
QUALITY CONTROL
1. Run quality control materials in normal and abnormal ranges (low and high
controls) at the beginning of each day. Always run a set of controls after
calibrating the analyser. If the controls are out of range, do not report patient
results until the problem is resolved and the patient samples are repeated with
quality control samples within acceptable range.
2. Variability is expressed as standard deviation (SD) and coefficient of variation
CV) and plotted on a Levy-Jennings graph.
Reference value:
Women up to 31 u/l at 37 ċ
Men up to 40 u/l at 37 ċ
RESULTS
1- Panic values: are double or triple to the normal value.
2- Becareful . Reference value for the end point procedure is differed from the
reference value for the kinetic procedure.
LIMITATIONS
A. High total protein samples that are predominantly gamma globulins can increase
ALT results. The sample should be diluted with an equal volume of isotonic
saline and then reanalysed.
B. In sera with very high activities, the initial absorbance may be very low because
most of the NADH may have been consumed before the first readings. In this
case, return the sample after dilution as described above.
8.2.2 Aspartate Aminotransferase (AST)
AST (Aspartate transaminase ): AST is normally found in red blood cells, liver, heart,
muscle tissue, pancreas, and kidneys. AST formerly was called serum glutamic
oxaloacetic transaminase.
Low levels of AST are normally found in the blood. When body tissue or an organ such
as the heart or liver is diseased or damaged, additional AST is released into the
bloodstream. The amount of AST in the blood is directly related to the extent of the tissue
damage. After severe damage, AST levels rise in 6 to 10 hours and remain high for about
4 days.
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The AST test may be done at the same time as a test for alanine aminotransferase, or
ALT. The ratio of AST to ALT sometimes can help determine whether the liver or
another organ has been damaged. Both ALT and AST levels can test for liver damage.
Causes of increase AST:
 Alcohol abuse .
 viral hepatitis,
 acute MI
Principle:
Kinetic determination of ALT activity , method recommended by IFCC
2-oxoglutarate +l-aspartate-----AST-------- glutamate + oxaloacetate
oxaloacetate + NADH + H+-----------MDH------- malate + NAD+
The rate of NADH consumption is determined photometrically and is direct proportional
to the ALT activity in the sample.
SPECIMEN
A. Recommended specimen: 100 μl of serum or heparinised plasma or EDTA
plasma. Collect specimens by standard venipuncture technique. Heparin may be
used as an anticoagulant for plasma specimens. Handle specimens in stoppered
containers to avoid contamination and evaporation. Follow universal precautions
when performing phlebotomy or handling patient specimens, calibrators, or other
serum-based products. Discard contaminated materials with infectious waste.
B. Remove serum promptly from the clot. Haemolysed specimens should not be used
because AST contamination from red blood cells will occur. Refrigerate
specimens if analysis is not performed immediately. Freeze specimens if analysis
is not performed within 48 hours.
C. If concentration is greater than the analyser range, dilute with an equal volume of
isotonic saline and reanalyse. Multiply the result by 2 to obtain the original AST
activity.
REAGENTS AND MATERIALS
1- full automated auto analyzer or spectrophotometer
2- Gel tube (clot activator)
3- Centrifuge
4- Reagents :
Reagent 1
tris buffer P H 7.8
80mmol/l
Buffer reagent
L-aspartate
200 mmol/l
Reagent 2
NADH
0.18 mmol/l
Substrate
LDH
800u/l
Oxoglutarate
12 mmol/l
MDH
600 u/l
Preparation:
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Dissolve one vial of reagent 2 with 3 ml of R 1 buffer , this working reagent stable 21
days at 2 – 8 ċ or 24 hours at 20 – 25 days .
Procedure:
 Wave length ----------340 nm
 Working reagent 1 ml mixed with sample 100 µl and wait 1 minute
 Measure the extinction decrease per minute for 1 – 3 minutes

Calculation:
Δ OD/min * 1750 = u/l
Reference value:
 Women up to 31 u/l at 37 ċ
 Men up to 38 u/l at 37 ċ
QUALITY CONTROL
A. Run quality control materials in normal and abnormal ranges (low and high
controls) at the beginning of each shift. Always run a set of controls after
calibrating the analyser. If the controls are out of range, do not report patient
results until the problem is resolved and the patient samples are repeated with
quality control samples within acceptable range.
B. Variability is expressed as standard deviation (SD) and coefficient of variation
(CV) and plotted on a Levy-Jennings graph.
LIMITATIONS
A. High total protein samples that are predominantly gamma globulins can increase
AST results. The sample should be diluted with an equal volume of isotonic saline
and reanalysed. Highly active sera can have a very low initial absorbance because
much of the NADH is already consumed prior to measurement. In such cases,
dilute sample as specified.
B. Analyser range: ≤ 150 U/L
8.2.3 Alkaline Phosphatase
Alkaline phosphatase is an enzyme used by the cells to speed up the time of a reaction. It
is found in the bone, placenta, kidneys, small intestines and liver, but is primarily
associated with liver disease and the obstruction of bile flow. Within the liver, alkaline
phosphatase is located near or within the tubes that collect bile.
Principle:
Phenyl phosphate-----------ALP-------- phenol + phosphate
Free phenol liberate by hydrolysis of substrate reacts then with 4-amino-antipyrine in the
presence of alkaline.
Potassium ferricyanide to form a red – colored complex which absorbance measured at
510 nm is directly proportional to ALP activity . sodium arsenate incorporated in the
reagent abolishes further enzyme activity and prevent the dilution of the color inherent in
earlier methods .
SPECIMEN
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Recommended specimen: 20 μl of serum or heparinised plasma. Collect specimens by
standard venipuncture technique. Handle specimens in stoppered containers to avoid
contamination and evaporation. Follow universal precautions when performing
phlebotomy or handling patient specimens, calibrators, or other serum-based products.
Discard contaminated materials with infectious waste.
Haemolysed specimens should not be used and samples more than 7 days old should not
be used due to loss of the enzyme activity in 7 days.
If absorbance change per minute exceeds 0.250, dilute 0.1 ml of the sample with 0.5 ml
of normal saline and repeat the assay using this dilution. Multiply the results by 6 to
obtain the alkaline phosphatase activity.
REAGENTS AND MATERIALS
A. full automated autoanalyzer or spectrophotometer .
B. Gel tube (clot activator)
C. Centrifuge
D. Reagents :
reagent 1
substrate buffer
disodium phenylphosphate
carbonate-biocarbonate buffer
P H 10 stabilizer
Reagent 2
phenol
Standard
Reagent 3
4- aminoantipyrine
Blocking reagent
sodium arsenate
Reagent 4
potassium ferricyanide
Dye reagent
5 mmol/l
50 mmol/l
20u/l
60 mmol/l
240 g/l
150 mmol/l
CALIBRATION
A. Empty and wash flow-through cuvette by pressing [WASH] on the LCD display.
An automatic return to the select menu will occur.
B. Push sip by pressing lever [P] to drain the tube system. Do not sip any solution.
C. Measure exactly 1,000 μl of distilled water into a sample cup and sip distilled
water by pushing sip-press lever [P]. The sipping volume of 1,000 μl is
automatically measured in the flow-through cuvette.
D. The length of transportation of the air and water will be displayed on the screen.
E. Press [↵] to finish calibration. The program will return to method select menu.
QUALITY CONTROL
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A. Run quality control materials in normal and abnormal ranges (low and high
controls) at the beginning of each shift. Always run a set of controls after
calibrating the analyser. If the controls are out of range, do not report patient
results until the problem is resolved and the patient samples are repeated with
quality control samples within acceptable range.
B. Variability is expressed as standard deviation (SD) and coefficient of variation
(CV) and plotted on a Levy-Jennings graph.
PROCEDURE
Serum blank
R1
2 ml
Incubate for 5 minutes at 37 ċ
Serum
R2
Incubate for 15 minutes at 37 ċ
R3
0.5 ml
Mix well
R4
0.5 ml
Serum
0.5µl
D.W.
-
Serum sample
2ml
Standard
2ml
Regent blank
2ml
50 µl
-
50 µl
-
0.5 ml
0.5 ml
0.5 ml
0.5 ml
-
0.5 ml
-
0.5 ml
50 µL
Mix well wait for 10 minutes in dark place, read at 510 nm against blank .
Calculation:
Conc.of ALP = O.D of serum / O.D. of standard * concentration of standard
Linearity: up to 285 u /l
Reference value:
children 17 – 142 u / l
Adult 21 – 92 u /l
Conv F.: kau/dl * 7.09 = u/l
8.2.4 Total Serum Bilirubin
bilirubin is formed from the haem fragment of haemoglobin released by damaged or aged
RBC . bone marrow , liver and spleen are the sites of bilirubin production .
Causes for an increase in total bilirubin may be divided into 3 categories:
 Pre- hepatic: resulting from various haemolytic states;
 Hepatic: resulting from hepatitis, cirrhosis, and other causes of hepatic necrosis;\
 Post-hepatic: resulting from an obstruction of the common bile or hepatic duct.
Principle:
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Sulfanilic acid react with sodium nitrate to form diazotized sulfanilic acid . in the
presence of dimethyl sulfoxide . total bilirubin react with dizotized sulfanilic acid form
azobilirubin .
SPECIMEN
A. Recommended specimen is 100 μl of serum or heparinised plasma. Collect
specimens by standard venipuncture technique. Handle specimens in stoppered
containers to avoid contamination and evaporation. Follow universal precautions
when performing phlebotomy or handling patient specimens, calibrators, or other
serum-based products. Discard contaminated materials with infectious waste.
B. Protect specimens from light and analyse as soon as possible after collection.
Direct exposure to sunlight is reported to cause as much as 50% loss of bilirubin
in one hour, especially when the specimen is kept in a capillary tube. Exposure to
normal room light can result in a significant loss of serum bilirubin after 2 to 3
hours.
C. If analysis is not performed immediately, samples can be refrigerated for up to 24
hours at (2–5)°C.
D. Samples that come in contact with alcohol from sterile wipes may become
haemolysed, which will increase the value of bilirubin.
E. If concentration is greater than the analyser range, dilute with 5 volumes of
normal saline and reanalyse. Multiply the result by 6 to obtain the original total
bilirubin concentration. The results may show a positive bias of up to 30%.
REAGENTS AND MATERIALS
A. spectrophotometer and bilirubimnometer for neonate .
B. Centrifuge
C. Gel tube (clot activator)
D. Reagents :
Reagent 1
sulfanilic acid
30 mmol/l
Hydrochloric acid
150 mmol/l
Dimethylsulfoxide
7 mol/l
Reagent 3
sodium nitrate
20 mmol/l
Reagent 4
Standard
Procedure:
Working reagent:
Mix
Reagent 1
20 volumes
Reagent 3
1 volume
Stability in absence of light
6 hours at 20-25 ċ
2 days at + 4 ċ
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Wave length -------------- 555nm
Temperature ------------- 37 ċ
Read against R1 sample blank
Read against R1 + R3 (working reagent ) sample and standard
CALIBRATION
A. Empty and wash flow-through cuvette by pressing [WASH] on the LCD display.
An automatic return to the select menu will occur.
B. Push sip by pressing lever [P] to drain the tube system. Do not sip any solution.
C. Measure exactly 1,000 μl of distilled water into a sample cup and sip distilled
water by pushing sip-press lever [P]. The sipping volume of 1,000 μl is
automatically measured in the flow-through cuvette.
D. The length of transportation of the air and water will be displayed on the screen.
E. Press [↵] to finish calibration. The program will return to method select menu.
QUALITY CONTROL
A. Run quality control materials in normal and abnormal ranges (low and high
controls) at the beginning of each shift. Always run a set of controls after
calibrating the analyser. If the controls are out of range, do not report patient
results until the problem is resolved and the patient samples are repeated with
quality control samples within acceptable range.
B. Variability is expressed as standard deviation (SD) and coefficient of variation
(CV) and plotted on a Levy-Jennings graph.
PROCEDURE
CALCULATIONS
Sample
Calibrator(R4)
Reagent(R1)
Working
reagent
Sample blank
50 µl
1 ml
-
Sample
50 µl
1 ml
Standard blank
50 µl
1 ml
-
Standard
50 µl
1 ml
Mix and read the extinction after incubation for 5 minutes at 37 ċ
Stability of color is 1 hour ( ovoid direct sunlight )
Calculation:
( O.D sample – O. D. sample blank ) / ( O. D. calibrator – O. D. calibrator blank ) * n
N = calibrator concentration
Linearity: linear up to 20 mg / dl , 340 µmol/l
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Reference value:
 Serum total bilirubin: < 1 mg/dl, < 17 µmol / l
 Conv. F. : mg / dl * 17.1 = µmol/l
LIMITATIONS
A. Specimens from haemodialysis patients should not be analysed for total bilirubin.
B. Haemoglobin affects bilirubin results.
C. Compounds that discolour serum, such as 4-aminosalicylic acid and
phenazopyridine, may falsely increase bilirubin results.
D. Results for predominantly unconjugated bilirubin (e.g., for neonates) may be up to
10% higher than the reference method.
8.3 Renal Function test
8.3.1 Blood Urea
The liver produces urea in the urea cycle as a waste product of the digestion of protein ,
Elevated levels of blood urea are most commonly associated with renal disease, but may
also result from dehydration, a high-protein diet, excess destruction of body proteins, and
gastrointestinal diseases, especially with intestinal obstruction.
Principle:
Urea + H2O --------urease------ CO2 + 2NH3
Salicylate and hypochlorite in the reagent react with the ammonium ions to form
2.2dicarboxy –indophenol . the quantity of green compound is proportional to the urea
concentration.
SPECIMEN
A. Recommended specimen is 10 μl of serum or EDTA plasma. Collect specimens
by standard venipuncture technique. Heparin may be used as an anticoagulant for
plasma specimens. Handle specimens in stoppered containers to avoid
contamination and evaporation. Follow universal precautions when performing
phlebotomy or handling patient specimens, calibrators, or other serum-based
products. Discard contaminated materials with infectious waste.
B. Refrigerate specimens up to 3 days at 4°C if analysis is not performed
immediately.
C. If concentration is greater than the analyser range, dilute with 9 volumes of
isotonic saline or distilled water and reanalyse. Multiply the result by 10 to obtain
the original urea nitrogen concentration.
REAGENTS AND MATERIALS
A. full automated auto analyzer or spectrophotometer
B. Gel tube (clot activator)
C. Centrifuge
D. Reagent :
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R1a
R1b
R2
CAL
urease
phosphate buffer
Sodium salicylate
Sodium nitroprusside
EDTA
sodium hypochloride
Sodium hydroxide
standard
>5000 u/l
120 mmol/l p H 7
63.4 mmol/l
5 mmol/l
1.5 mmol/l
18 mmol/l
750 mmol/l
8.05 mmol/l
Preparation:
Add 1 vial of of urease R1a to one bottle of phosphate buffer R1b
Procedure :
Wave length …….600 nm
Temperature ……25 ċ – 37 ċ
CALIBRATION
A. Empty and wash flow-through cuvette by pressing [WASH] on the LCD display.
An automatic return to the select menu will occur.
B. Push sip by pressing lever [P] to drain the tube system. Do not sip any solution.
C. Measure exactly 1,000 μl of distilled water into a sample cup and sip distilled
water by pushing sip-press lever [P]. The sipping volume of 1,000 μl is
automatically measured in the flow-through cuvette.
D. The length of transportation of the air and water will be displayed on the screen.
E. Press [↵] to finish calibration. The program will return to method select menu.
QUALITY CONTROL
A. Run quality control materials in normal and abnormal ranges (low and high
controls) at the beginning of each shift. Always run a set of controls after
calibrating the analyser. If the controls are out of range, do not report patient
results until the problem is resolved and the patient samples are repeated with
quality control samples within acceptable range.
B. Variability is expressed as standard deviation (SD) and coefficient of variation
(CV) and plotted on a Levy-Jennings graph.
PROCEDURE
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Blank
Standard
standard
10µl
sample
Working reagent
1 ml
1 ml
Mixture incubated for at least 3 minutes at 37 ċ or 5 min. at 20-25 ċ
sample
10µl
1 ml
Sodium
200 µl
200 µl
200 µl
hypochloride
mix. incubate for at least 5 min. at 37 ċ or 10 min. at 20-25 ċ , measurement is against
reagent blank .
Calculation:
O.D.sample / O.D. standard * concentration of standard
CALCULATIONS
The results are reported in mmol/L. No further calculation is necessary unless the
specimen has been diluted.
Conversion
Urea = 2.14 × BUN
BUN = 0.466 × Urea
RESULTS
A. Expected values: (urea) 1.7–8.3 mmol/L
B. Panic values: <1.07 mmol/L and >37.1 mmol/L
C. Each laboratory should confirm these values.
LIMITATIONS
A. Do not use plasma collected with sodium fluoride.
B. Specimens that contain haemoglobin increase urea nitrogen. Haemoglobin of 50
mg/dL (slightly haemolysed) increases BUN levels below 28 mg/dL by 1 mg/dL.
C. Ammonium ions have shown an increase in urea nitrogen equivalent to their
nitrogen content.
8.3.2 Serum Creatinine
Creatinine measurement is used in the diagnosis and treatment of renal diseases and in
monitoring patients on renal dialysis.
Analysis of creatinine activity in 100 μl of serum or plasma is based on an enzymatic
method that produces a coloured product.
A. The 100 μl of patient sample mixed with picrate in alkaline medium produces a
coloured complex with creatinine. The rate of formation of the complex is then
measured.
252 | P a g e
B. The creatinine present in the sample reacts with the picrate at 37°C, and the rate
of complex formation is measured and is proportional to the creatinine
concentration.
SPECIMEN
A. Recommended specimen is 100 μl of serum or heparinized plasma. Collect
specimens by standard venipuncture technique. Heparin may be used as an
anticoagulant for plasma specimens. Handle specimens in stoppered containers to
avoid contamination and evaporation. Follow universal precautions when
performing phlebotomy or handling patient specimens, calibrators, or other
serum-based products. Discard contaminated infectious waste.
B. Refrigerate specimens at 4°C for 24 hours if analysis is not done immediately.
REAGENTS AND MATERIALS
A. Photometer
B. Gel tube (clot activator)
C. Centrifuge
D. Picric acid 26 mmol/L
E. NaOH 1.6 mmol/L
F. Creatinine 2 mg/dL or 176.8 μmol/L
CALIBRATION
A. Empty and wash flow-through cuvette by pressing [WASH] on the LCD display.
An automatic return to the select menu will occur.
B. Push sip by pressing lever [P] to drain the tube system. Do not sip any solution.
C. Measure exactly 1,000 μl of distilled water into a sample cup and sip distilled
water by pushing sip-press lever [P]. The sipping volume of 1,000 μl is
automatically measured in the flow-through cuvette.
D. The length of transportation of the air and water will be displayed on the screen.
E. Press [↵] to finish calibration. The program will return to method select menu.
QUALITY CONTROL
A. Run quality control materials in normal and abnormal ranges (low and high
controls) at the beginning of each shift. Always run a set of controls after
calibrating the analyser. If the controls are out of range, do not report patient
results until the problem is resolved and the patient samples are repeated with
quality control samples within acceptable range.
B. Variability is expressed as standard deviation (SD) and coefficient of variation
(CV) and plotted on a Levy-Jennings graph.
PROCEDURE
A. Mix reagents according to manufacturer’s recommendations.
253 | P a g e
B. Label 2 tubes: in tube or standard tube pipette 1,000 μl of reagents solution and
100 μl of creatinine standard reagent; and in tube 2 or sample tube pipette 1,000
μl of reagents solution and 100 μl of sample.
C. Mix them well and measure them immediately as indicated.
D. Press ZERO and then push sip by pressing lever [P] to sip water and drain the
system.
E. When the screen displays MEASURE STANDARD, place cuvette with the
standard solution into cuvette holder and aspirate the standard solution by pushing
lever [P].
F. When the screen displays MEASURE SAMPLE, place cuvette with the sample
solution into cuvette holder and aspirate the sample solution by pushing lever [P].
G. Press [RESULT] and await possible delay while measuring the sample solution.
The result will be displayed on top line.
H. Print or write out the results. Press [WASH] to sip rinse solution or press
[MEASURE] to measure the next sample by pushing sip lever [P].
CALCULATIONS
The results are reported in μmol/L. No further calculation is necessary unless the
specimen has been diluted.
RESULTS
A. Expected values:
 Males: 53–97 μmol/L
 Females: 44–80 μmol/L
B. Panic values: < 27 μmol/L and > 663 μmol/L
C. Each laboratory should confirm these values.
LIMITATIONS
A. EDTA and fluoride/oxalate anticoagulants will cause low creatinine results.
B. Lidocaine: Patients on long-term lidocaine therapy may show an increase of up to
1.0 mg/dL. Ninety percent of patients receiving intravenous lidocaine will show
less than a 0.3 mg/dL increase.
C. Proline: Patients receiving hyperalimentation fluid that contains proline may show
an increase of up to 2.0 mg/dL.
D. TRIS buffer (control fluid) causes an approximate 50% decrease in results.
E. Dipyrone (Metamizol) at 40 mg/dL shows a –0.6 mg/dL bias at a creatinine
concentration of 1.0 mg/dL.
F. N-acetylcysteine: Patients receiving N-acetylcysteine (Fluimucil, Mucomyst)
intravenously have been reported to show a large negative bias.
8.3.3 Uric acid
Uric acid is a chemical created when the body breaks down substances called purines.
Purines are found in some foods and drinks ,Most uric acid dissolves in blood and
254 | P a g e
travels to the kidneys, where it passes out in urine. High levels of uric acid in the
body is called hyperuricemia.
Principle :
Uric acid +O2 +H2O---------uricase -------- allantoine + CO2 + H2O2
2H2O2 +4-aminophenazone+ 2-4dichloro- phenol-sulfonate -----------peroxidase------- quinonemine
Reagents:
Reagent 1
phosphate buffer P H 7.4
50mmol/l
Buffer solution
2-4 DCPS
4 mmol/l
Reagent 2
uricase
7o u/l
Vial of enzyme
peroxidase
660 u/l
4-aminophenazone
1 mmol/l
uric acid
6 mg/dl
Reagent 3
Standard
Preparation :
Dissolve the content of one bottle R 2 with the content of one bottle buffer R 1
Procedure :
Wave length---------------------510 nm (490 – 550)
Blank
Standard
Sample
Working reagent
1 ml
1 ml
1 ml
Standard
-
20µl
-
Sample
-
-
20 µl
mix. incubate 5 minutes at 37 ċ or 10 minutes at 20-25 ċ the colour is stable for 30
minutes .
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calculation :
concentration= O.D.sample / O.D. standard * concentration of standard
reference value : women 2.5 – 6 mg/dl,148 – 357 µmol / l
men 3.4 – 7 mg/dl , 200 – 416 µmol / l
Conv. F. : mg/dl * 60 = µmol / l
8.4 Lipid Profile
8.4.1 Total Cholesterol
Cholesterol is a waxy, fat-like substance that occurs naturally in all parts of the body.
Your body needs some cholesterol to work properly. But if you have too much in your
blood, it can stick to the walls of your arteries. This is called plaque. Plaque can narrow
your arteries or even block them.
High levels of cholesterol in the blood can increase your risk of heart disease. Your
cholesterol levels tend to rise as you get older. There are usually no signs or symptoms
that you have high blood cholesterol, but it can be detected with a blood test. You are
likely to have high cholesterol if members of your family have it, if you are overweight or
if you eat a lot of fatty foods.
You can lower your cholesterol by exercising more and eating more fruits and vegetables.
You also may need to take medicine to lower your cholesterol.
Causes of hypercholesterolemia:
1- familial hypercholesterolaemia (FH)
2- reduced metabolism due to thyroid problems(hypothyroidism)
3-kidney diseases
4- diabetes particularly when poorly controlled
5- alcohol abuse
6- being overweight – this is probably the commonest cause of high cholesterol levels.
Causes of hypocholesterolemia :
1-Hyperthyroidism
2-Liver disease
3-Malnutrition
4-Cancer
5-Chronic infections or inflammation
Principle:
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Esters of cholesterol +H2O -------chloesterol esterase-------> cholesterol + fatty acids
Cholesterol +O2 ---------cholesterol oxidase -----------> cholest-4-en-one +H2O2
H2O2 +phenol+4-aminophenazone ------------peroxidase ------------> quinonimine
SPECIMEN
A. Recommended specimen 10 μl of serum, heparinised or EDTA plasma. Collect
specimens by standard venipuncture technique. Heparin may be used as an
anticoagulant for plasma specimens. Handle specimens in stoppered containers to
avoid contamination and evaporation. Follow universal precautions when
performing phlebotomy or handling patient specimens, calibrators, or other
serum-based products. Discard contaminated materials with infectious waste.
B. If the result is over 19.3 mmol/L, dilute the sample before precipitation 1:2 with
normal saline and multiply the results by 3 to obtain the original cholesterol
concentration.
C. Cholesterol remains stable in serum for 6 days at 4–25°C and up to 4 months at –
20°C.
D. Lipemic specimens generate turbidity of the sample/reagent mixture, which leads
to falsely elevated results.
REAGENTS AND MATERIALS
A. spectrophotometer
B. Gel tube (clot activator)
C. Centrifuge
D. Reagents :
Reagent 1
pipes p H 6.9
90 mmol/l
Buffer solution
Phenol
26 mmol/l
Reagent 2
cholesterol oxidase
399 u/l
Vial of two enzymes
Peroxidase
1250 u/l
Cholesterol oxidase
300 u/l
4-aminophenazone
0.4 mmol/l
Reagent 3
standard
200 mg/dl
Preparation:

Dissolve the contents of one bottle R2 to the contents of one bottle buffer reagent
R1
Sample: serum , heparinized plasma
Procedure:


Wave length …………… 505nm ( 500 550)
Temperature …………... 37 ċ
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Blank
Standard
standard
10µl
sample
Working reagent
1 ml
1 ml
 Mix. incubate 5 minutes at 37 ċ ,the colour is stable for 30 min.
Calculation:
sample
10µl
1 ml
Cholesterol concentration = O.D.sample / O.D. standard * concentration of standard
QUALITY CONTROL
A. Run quality control materials in normal and abnormal ranges (low and high
controls) at the beginning of each shift. Always run a set of controls after
calibrating the analyser. If the controls are out of range, do not report patient
results until the problem is resolved and the patient samples are repeated with
quality control samples within acceptable range.
B. Variability is expressed as standard deviation (SD) and coefficient of variation
(CV) and plotted on a Levy-Jennings graph.
CALCULATIONS
The results are reported in mmol/L. No further calculation is necessary unless the
specimen has been diluted.
RESULTS
A. Expected values: 3.6–5.2 mmol/L.
B. Each laboratory should confirm these values.
LIMITATIONS
A. Bilirubin concentrations above 4 mg/dl interferes with tests
B. If results are over 25.9mmol/l dilute the sample and repeat measument
C. High concentration of ascorbic acid (> 2.5 mg/dL) will give lower values.
D. Haemoglobin levels higher than 200 mg/dL do not interfere with the test.
8.4.2 Triglycerides
Triglycerides protein is essential for growth, the production of new tissue, and the repair
of injured tissue. An increase in triglycerides levels may be the result of nephrosis,
cholestasis, pancreatitis, cirrhosis, diabetes mellitus, and hepatitis. Adecrease is seen with
malnutrition.
The analysis is based on the enzymatic hydrolysis of triglycerides with lipases in a
quinoneimine formed from hydrogen peroxide used as a coloured indicator. The intensity
of the colour is proportional to the amount of triglycerides in the sample. 10 μl of patient
sample is mixed and incubated with 1,000 μl of solution containing 4- aminoantipyrine
and 4-chlorophenol in presence of hydrogen peroxide. After incubation for 10 minutes, a
reaction between 4-aminoantipyrine and 4-chlorophenol and hydrogen peroxide takes
place, forming quinoneimine, which is a coloured
258 | P a g e
product. The amount of coloured compound formed is proportional to the amount of
triglycerides in the specimen.
SPECIMEN
A. Recommended specimen 10 μl of serum, heparinised or EDTA plasma. Collect
specimens by standard venipuncture technique. Heparin may be used as an
anticoagulant for plasma specimens. Handle specimens in stoppered containers to
avoid contamination and evaporation. Follow universal precautions when
performing phlebotomy or handling patient specimens, calibrators, or other
serum-based products. Discard contaminated materials with infectious waste.
B. Avoid haemolysed specimens. If analysis is not performed immediately, freeze
specimens at −25oC for up to 4 months or refrigerate at 2–8oC for up to 3 days.
C. If concentration is greater than the analyser range, dilute 1 part of sample with 4
parts of isotonic saline or distilled water and reanalyse. Multiply the result by 5 to
obtain the original triglycerides concentration.
REAGENTS AND MATERIALS
A. Photometer
B. Gel tube (clot activator)
C. Centrifuge
D. Reagent PIPES buffer (pH 7.5)
50 mmol/L
 4-chlorophenol
5 mmol/L
 4-aminoantipyrine
0.25 mmol/L
 Magnesium ion
4.5 mmol/L
 ATP
2 mmol/L
 Lipases
≥1.3 U/ml
 Peroxidase
≥0.5 U/ml
 Glycerol kinase
≥0.4 U/ml
 Glycerol-3-phpsphate oxidase
≥1.5 U/ml
 Standard Triglyceride
2.28 mmol/L
CALIBRATION
A. Empty and wash flow-through cuvette by pressing [WASH] on the LCD display.
An automatic return to the select menu will occur.
B. Push sip by pressing lever [P] to drain the tube system. Do not sip any solution.
C. Measure exactly 1,000 μl of distilled water into a sample cup and sip distilled
water by pushing sip-press lever [P]. The sipping volume of 1,000 μl is
automatically measured in the flow-through cuvette.
D. The length of transportation of the air and water will be displayed on the screen.
E. Press [↵] to finish calibration. The program will return to method select menu.
QUALITY CONTROL
259 | P a g e
A. Run quality control materials in normal and abnormal ranges (low and
highcontrols) at the beginning of each shift. Always run a set of controls after
calibrating the analyser. If the controls are out of range, do not report patient
results until the problem is resolved and the patient samples are repeated with
quality control samples within acceptable range.
B. Variability is expressed as standard deviation (SD) and coefficient of variation
(CV) and plotted on a Levy-Jennings graph.
PROCEDURE
A. Mix 2 tubes: one blank with 1,000 μl of standard reagent, and tube 2 with 10 μl of
serum or plasma and 1,000 μl of reagent solution.
B. Mix each well and incubate for 5 minutes at 37oC, and measure the absorbance of
the standard reagent and sample immediately as indicated.
C. Press ZERO and then push sip by pressing lever [P] to sip water and drain the
system.
D. When the screen displays MEASURE RB, place cuvette with the blank solution
into cuvette holder and aspirate the blank solution by pushing lever [P].
E. When the screen displays MEASURE SAMPLE, place cuvette with the sample
solution into cuvette holder and aspirate the sample solution by pushing lever [P].
F. Press [RESULT] and await possible delay while measuring the sample solution.
The result will be displayed on top line.
G. Print or write out the results. Press [WASH] to sip rinse solution or press
[MEASURE] to measure the next sample by pushing sip lever [P].
CALCULATIONS
The results are reported in mmol/L. No further calculation is necessary unless the
specimen has been diluted.
RESULTS
A. Expected values: 0.11–2.15 mmol/L.
B. Each laboratory should confirm these values.
LIMITATIONS
A. Lipemic specimen usually generates turbidity of the sample reagent mixture,
which leads to falsely elevated results.
B. Ascorbate gives falsely low values.
8.4.3 HDL Cholesterol
High density lipoprotein (HDL) is lipoprotein which is involved in the transport of
cholesterol from the cells to the liver. An increase in HDL levels may be the result of
nephrosis, cholestasis, pancreatitis, cirrhosis, diabetes mellitus, and hepatitis. A decrease
is seen with malnutrition.
The analysis is based on the precipitation of chylomicrons, very low-density lipoprotein
(VLDL), and low density lipoprotein (LDL) upon addition of phospshotungstic acid and
260 | P a g e
magnesium chloride to the sample. The HDL is contained in the supernatant, which can
be enzymatically assayed after centrifugation of the supernatant. The absorbance of the
sample and the standard reagent is measured against blank reagent.
When 200 μl of test sample is mixed with 500 μl of precipitant reagent containing
phosphotungstic acid and magnesium chloride at room temperature for 10 minutes
chylomicrons, VLDL, and LDL are precipitated . Centrifugation at 10,000 g for 2
minutes leaves only the HDL in the supernatant, where the concentration of HDL in the
sample is determined enzymatically.
SPECIMEN
A. Recommended specimen 200 μl of serum, heparinised or EDTA plasma. Collect
specimens by standard venipuncture technique. Heparin may be used as an
anticoagulant for plasma specimens. Handle specimens in stoppered containers
toavoid contamination and evaporation. Follow universal precautions when
performing phlebotomy or handling patient specimens, calibrators, or otherbased products. Discard contaminated materials with infectious waste.
B. If the supernatant is not clear, dilute the sample before precipitation 1:1 with
normal saline and multiply the results by 2 to obtain the original HDL
concentration.
C. HDL cholesterol remains stable in serum for 6 days at 4–25°C and up to 4 months
at –20°C. Serum must be separated from the blood clot as rapidly as possible.
REAGENTS AND MATERIALS
A. Photometer
B. Gel tube (clot activator)
C. Centrifuge
D. Reagent (Precipitant)
Phosphotungstic acid
0.55 mmol/L
Magnesium chloride
25 mmol/L
Standard Cholesterol
1.29 mmol/L
CALIBRATION
A. Empty and wash flow-through cuvette by pressing [WASH] on the LCD display.
An automatic return to the select menu will occur.
B. Push sip by pressing lever [P] to drain the tube system. Do not sip any solution.
C. Measure exactly 1,000 μl of distilled water into a sample cup and sip distilled
water by pushing sip-press lever “P”. The sipping volume of 1,000 μl is
automatically measured in the flow-through cuvette.
D. The length of transportation of the air and water will be displayed on the screen.
E. Press [↵] to finish calibration. The program will return to method select menu.
QUALITY CONTROL
261 | P a g e
A. Run quality control materials in normal and abnormal ranges (low and high
controls) at the beginning of each shift. Always run a set of controls after
calibrating the analyser. If the controls are out of range, do not report patient
results until the problem is resolved and the patient samples are repeated with
quality control samples within acceptable range.
B. Variability is expressed as standard deviation (SD) and coefficient of variation
(CV) and plotted on a Levy-Jennings graph.
PROCEDURE
A. Into a glass centrifuge tube mix 200 μl of test sample (serum or plasma) 500 μl of
precipitant reagent. (Dilute precipitant reagent with distilled water 4 in 1.)
B. Mix them well and incubate for 10 minutes at room temperature.
C. Centrifuge at 4000 rpm for 10 minutes or 10,000 rpm for 2 minutes.
D. After centrifugation remove the clear supernatant and into tube 1 (sample) add
100 μl of supernatant and 1,000 μl of total cholesterol reagent. In tube 2 (reagent
blank) add 100 μl of distilled and 1,000 μl of cholesterol reagent.
E. Mix them well and incubate for 10 minutes at 20–25°C or 5 minutes at 37°C and
measure their absorbance.
F. When the screen displays MEASURE BLANK, press ZERO and then push sip by
pressing lever [P] to sip water and drain the system.
G. When the screen displays MEASURE RB, place cuvette with the blank solution
into cuvette holder and aspirate the blank solution by pushing lever [P].
H. When the screen displays MEASURE SAMPLE, place cuvette with the sample
solution into cuvette holder and aspirate the sample solution by pushing lever [P].
I. Press [RESULT] and await possible delay while measuring the sample solution.
The result will be displayed on top line.
J. Print or write out the results. Press [WASH] to sip rinse solution or press
[measure] to measure the next sample by pushing sip lever [P].
CALCULATIONS
The results are reported in mmol/L. No further calculation is necessary unless the
specimen has been diluted.
RESULTS
A. Expected values: 0–0.9 mmol/L.
B. Each laboratory should confirm these values.
LIMITATIONS
A. High triglyceride level interferes with HDL measure; dilute the sample before
precipitation 1:1 with 0.9% saline and multiply the results by 2.
B. High concentration of ascorbic acid (> 2.5 mg/dL) will give lower values.
C. Haemoglobin levels higher than 100 mg/dL and bilirubin levels higher than 10
mg/dL interfere with the test.
262 | P a g e
8.4.4 LDL Cholesterol
Low density lipoprotein (LDL) is lipoprotein which is involved in the transport of
cholesterol to the cells. An increase in LDL levels may be the result of altered diet and
vascular diseases. A decrease is seen in AIDS, haematological malignancies such as acute
myeloid leukaemia (AML) or chronic myeloid leukaemia (CML), and disorders involving
splenomegaly.
The analysis is based on direct homogenous enzymatic assay for quantitative
determination of LDL cholesterol combining two steps. The analysis involves removal of
chylomicrons, very low density lipoprotein (VLDL), and high density lipoprotein (HDL),
followed by enzymatic determination of LDL in the sample.
10 μl of patient sample is mixed and incubated at 37oC for 5 minutes with 750 μl of
enzyme solution containing cholesterol esterase and cholesterol oxidase, which allows the
removal of chylomicrons, VLDL, and HDL from the specimen by formation of
cholestenone and hydrogen peroxide. The LDL present in the sample is converted to
cholestenone and hydrogen peroxide in the presence of cholesterol esterase, cholesterol
oxidase, and specific surfactant. The chromogen in presence of hydrogen peroxide
peroxidase is converted to a coloured dye (quinine) whose concentration is measured by
colorimetric method.
SPECIMEN
A. Recommended specimen 10 μl of serum or plasma. Collect specimens by standard
venipuncture technique. Heparin may be used as an anticoagulant for plasma
specimens. Handle specimens in stoppered containers to avoid contamination and
evaporation. Follow universal precautions when performing phlebotomy or
handling patient specimens, calibrators, or other serum-based products. Discard
contaminated materials with infectious waste.
B. If the serum concentration of LDL exceeds the measuring range, dilute the sample
before precipitation 1:1 with normal saline, repeat the test, and multiply the
results by 2 to obtain the original LDL concentration.
C. LDL cholesterol remains stable in serum for 6 days at 4–25°C and up to 4 months
at –20°C.
D. Serum must be separated from the blood clot as rapidly as possible.
REAGENTS AND MATERIALS
A. Photometer
B. Gel tube (clot activator)
C. Centrifuge
D. Reagent (enzymes)
 Good’s buffer (pH 7.0)
50 mmol/L
 Cholesterol esterase
600 U/L
 Cholesterol esterase
500 U/L
263 | P a g e
 Catalase
600 kU/L
 TOOS
2.0 mmol/L
 Detergents
0.3% w/v
 Preservatives
<0.1% w/v
 Substrate Peroxidase
4000 U/L
 4-Aminoantipyrin
4 mmol/L
 Good’s buffer (pH 7.0)
50 mmol/L
 Sodium azide
0.05%
 Detergents
1% w/v
 Preservatives
<0.1% w/v
CALIBRATION
A. Empty and wash flow-through cuvette by pressing [WASH] on the LCD display.
An automatic return to the select menu will occur.
B. Push sip by pressing lever [P] to drain the tube system. Do not sip any solution.
C. Measure exactly 1,000 μl of distilled water into a sample cup and sip distilled
water by pushing sip-press lever [P]. The sipping volume of 1,000 μl is
automatically measured in the flow-through cuvette. The length of transportation
of the air and water will be displayed on the screen.
D. Press [↵] to finish calibration. The program will return to method select menu.
QUALITY CONTROL
A. Run quality control materials in normal and abnormal ranges (low and high
controls) at the beginning of each shift. Always run a set of controls after
calibrating the analyser. If the controls are out of range, do not report patient
results until the problem is resolved and the patient samples are repeated with
quality control samples within acceptable range.
B. Variability is expressed as standard deviation (SD) and coefficient of variation
(CV) and plotted on a Levy-Jennings graph.
PROCEDURE
A. Label 2 tubes and pipette 10 μl water and 750 μl of enzyme solution into tube 1
(blank), and into tube 2 pipette 10 μl of test sample and 750 μl of enzyme
solution. Mix gently and incubate at 37oC for 5 minutes.
B. Into each tube add 250 μl of substrate solution and mix them well and incubate for
5 minutes at 37°C, then measure absorbance of sample against reagent blank.
C. When the screen displays MEASURE BLANK, press ZERO and then push sip by
pressing lever [P] to sip water and drain the system.
D. When the screen displays MEASURE RB, place cuvette with the blank solution
into cuvette holder and aspirate the blank solution by pushing lever [P].
E. When the screen displays MEASURE SAMPLE, place cuvette with the sample
solution into cuvette holder and aspirate the sample solution by pushing lever [P].
264 | P a g e
F. Press [RESULT] and await possible delay while measuring the sample solution.
The result will be displayed on top line.
G. Print or write out the results. Press [WASH] to sip rinse solution or press
[MEASURE] to measure the next sample by pushing sip lever [P].
CALCULATIONS
The results are reported in mmol/L. No further calculation is necessary unless the
specimen has been diluted.
RESULTS
A. Expected values: 0–0.9 mmol/L.
B. Each laboratory should confirm these values.
NOTE: Direct measurement of LDL is required instead of the Friedwald, or “calculated”,
method whenever triglyceride values exceed 400 mg/dL. The Friedwald or “calculated”
method of LDL determination is determined by subtracting HDL and VLDL from the
total cholesterol concentration. Total cholesterol = HDL + LDL + VLDL. VLDL is
generally estimated by the triglyceride concentration divided by 5, but is inaccurate when
triglyceride values exceed 400 mg/dL.
LIMITATIONS
A. High triglyceride level interferes with LDL measure; dilute the sample before
precipitation 1:1 with 0.9% saline and multiply the results by 2.
B. High concentration of ascorbic acid (> 2.5 mg/dL) will give lower values.
C. Haemoglobin levels higher than 100 mg/dL and bilirubin levels higher than 10
mg/dL interfere with the test.
265 | P a g e
Annex 1
TABLE 1 - Outline of the properties of heat decontamination methods. For everyday
laboratory purposes, autoclaving is the preferred method, unless the item cannot
withstand the heat and/or moisture of autoclaving.
Principle/Conditions Advantages Disadvantages
Dry Heat
Thermal inactivation: Nondestroys by oxidation corrosive
Simple
design and
principle
Hot Air Oven · 160-180?C for 2-4
hours
· penetrates
waterinsoluble
materials
(e.g., grease
and oil)
Uses
Less effective
than moist
heat; requires
longer times
and/or higher
temperatures
Materials that
are damaged by,
or are
impenetrable to,
moist heat
· slow
diffusion,
penetration
· anhydrous
materials, such
as oils, greases
and powders
· loading,
packing critical · laboratory
to performance glassware,
· less
instruments
·
not
suitable
corrosive to
· closed
metals and for reusable
plastics
containers
sharp
instruments
than steam
Red-heat
Flame
· oxidation to ashes
(burning)
· rapid
· initial
contact with
flame can
· inoculating
loops, needles
266 | P a g e
produce a
viable aerosol
· possibility of
accidental fire
Incineration
· oxidation to ashes
(burning)
· 1-60 minutes:
temperatures may
exceed 1000?C
· reduces
volume of
waste by up
to 95%
· improper use
may lead to
emission of
pathogens in
smoke
· for
decontamination
of waste items
prior to disposal
in landfill
· requires
transport of
infectious
waste
· excess
plastic (>20%)
content
reduces
combustibility
Moist Heat
Irreversible
coagulation of
(microbial) proteins
Pasteurization · heating to below
boiling point
(generally 77?C) for
up to 30 minutes
More rapid
and more
effective
than dry
heat
· can be
· not reliably
used on heat sporicidal
sensitive
liquids and
medical
devices
· milk and dairy
products
· some heatsensitive
medical
equipment
· low cost
267 | P a g e
Tyndallization · heating to 80(Fractional
100?C for 30 mins on
Sterilization) successive days, with
incubation periods in
between
Boiling
autoclaving
· resistant
spores
germinate
and are
killed on the
second and
third days
· time
consuming
· not reliably
sporicidal
· heat sensitive
materials such
as bacteriologic
media, solutions
of chemicals,
biological
materials
· maximum
· minimal
temperature
equipment
obtainable is
required
approximately 100?C
10-30 mins
· cumbersome: · small
not practical
instruments and
for everyday
equipment
lab use
· steam under
pressure
· loading and · penetration of
packing critical sterile
to performance glassware,
media and
· shielding dirt instruments
must first be
removed
·
decontamination
· maintenance of reusable
and quality
supplies and
control
equipment
essential
·
· damages
decontamination
heat-sensitive of infectious
itmes
waste
· 121?C/15 psi for
15-90 mins (gravity
displacement
autoclave)
· 132?C/27 psi for 420 minutes (prevacuum autoclave)
· minimal
time
required
· most
dependable
sterilant for
lab use
· not reliably
sporicidal
TABLE 2 - Summary of concentrations used, contact times, advantages and
disadvantages and uses of some of the halogen-releasing chemical germicides. The wide
ranges of effective concentrations and contact times cited are due to a number of
factors, including the interdependence of time and concentration, the variability in
268 | P a g e
resistance of different microorganisms, the amount of organic material present and the
desired effect (e.g., low-level vs high-level disinfection)
Effective
Concentrations
Disadvantage
Advantages
, Contact
s
Times
Chlorine
Compounds:
Sodium hypochlorite
solution 1 (liquid
bleach)
· 100-10,000
ppm (.01-1%)
free chlorine
· broad
spectrum
·
· 10-60 minutes inexpensive
(>= 3,000 ppm
· widely
for broad
available
spectrum)
·
bactericidal
at low
temperature
Examples of
Uses
· toxic,
corrosive to
skin and
metals
· general
disinfectant
· unstable at
optimum
effective pH
of 6
· surface
decontaminatio
n
· inactivated
by organic
matter
· waste liquids
· emergency
spill clean up
· instrument
disinfection
· deteriorates
under light
and heat: shelf
life of
dilutions is
less than 1
week
Calcium
hypochlorite2
granules, powder,
tablets
· as for liquid
bleach
NaDCC3 (Sodium
· as for liquid
dichloroisocyanurate bleach
) powder, granules,
tablets
· as for
liquid bleach
but more
stable
· as for liquid · as for liquid
bleach above, bleach
except shelf
life is longer
· more
· toxic,
stable than
corrosive
hypochlorite
s
· as for liquid
bleach
269 | P a g e
Chloramine-T4
(Sodium
tosylchloramide)
powder or tablets
· as for liquid
bleach
· stable at
pH 6.0
· inactivated
by organic
matter
· more
stable, less
affected by
organic
matter than
hypochlorite
s
· deteriorates · as for liquid
under
bleach
humidity, light
and heat
· longer
activity than
hypochlorite
s
Chlorine dioxide5
· demandrelease of
chlorine dioxide
in situ
· longer
activity than
other
chlorine
compounds
· aqueous
solutions
decompose
under light
· instrument
disinfection
· not
consistently
sporicidal
· germicidal
soaps and
antiseptics
· less
corrosive,
less toxic
than other
chlorine
compounds
· gas
sterilization of
germ-free
animal
chambers
· effective
at pH 6-10
Iodine
Preparations:
Iodophors6
· 30-1,000 ppm · broad
(.003-.1%) free spectrum
iodine
· germicidal
· 10-30 minutes over a wide
pH range
· efficacy
· surface
reduced by
decontaminatio
organic matter n
270 | P a g e
· generally
nonstaining,
less toxic
and less
irritating
than aqueous
or alcoholic
iodine
solutions
· some
iodophor
solutions
support
growth of
Pseudomonas7
· work surface
wipedown
· instrument
disinfection
TABLE 3A Summary of recommended concentrations, contact times, advantages and
disadvantages of non-halogen chemical germicides. The wide ranges of effective
concentrations and contact times cited reflect the interdependence of time and
concentration as well as factors such as resistance of the particular class or strain of
target microorganism(s) and desired effect. Also, some germicides are available in
combinations (e.g., glutaraldehyde/phenol or peracetic acid/alcohol mixtures) which are
synergistic whereby the components in combination produce a greater antimicrobial
effect than the sum of their individual effects.
Effective
Concentrations
and Contact
Times
Alcohols
· 70-80%
ethanol
· 60-95%
isopropanol
· 10-30
minutes
Advantages
· low toxicity
· rapid action
· low residue
Disadvantages
· rapid
evaporation
limits contact
time
· non-corrosive · flammable,
eye irratant
Examples of
Laboratory
Uses
· skin
disinfectant
(antiseptic)
· surface
decontamination
· benchtop,
· may damage cabinet
rubber, plastic, wipedown
shellac
· ineffective
against
271 | P a g e
bacterial
spores
Phenolic
Compounds
· 400-50,000
· tolerant of
ppm (.05-1.5%) organic load,
"hard" dilution
· 10-30
water
minutes
· leaves an
active residue
(may be
desirable on
some surfaces)
· pungent
odour,
corrosive,
some forms
toxic
· instruments
and equipment
disinfection
· disinfection of
floors and other
surfaces
· not
sporicidal;
limited activity · antiseptic
against viruses soaps and
lotions
· biodegradable · leaves a
residual film
(undesirable in
culture
systems)
· may support
growth of
bacteria1
Quaternary
Ammonium
Compounds
· 500-15,000
· combined
ppm (.05-1.5%) detergent and
germicidal
· 10-30
activity
minutes
· stable
· working
dilutions have
low toxicity
· non
sporicidal,
limited activity
against viruses,
mycobacteria
· surface
decontamination
· most
formulations
not readily
biodegradable
· antiseptic
formulations
available
· equipment
wipedown
· floors and
· may support walls
growth of
bacteria2
272 | P a g e
Hydrogen
Peroxide
· 3-30%
aqueous
solution
· rapid action
· 10-60
minutes
· low toxicity
· 6% for 30
minutes may
kill spores
· no residue
· limited
sporicidal
activity
· corrosive to
some metals
· surface
decontamination
· instruments
and equipment
·
environmentally · potentially
safe
explosive at
high
concentrations
· stock
solutions
irritating to
skin and eyes
Peracetic Acid · .001-.3%
aqueous
(PAA)
solution
· broad
spectrum
· pungent
odour
· gas phase: 24%
· sporicidal at
low
temperatures
corrosive to
some metals
· 5-120
minutes
· can tolerate
organic load
· rapid action
· nontoxic
decomposition
products
· leaves no
residue
· instruments
and equipment
· gas phase
sterilization of
chambers for
· shelf life of germ-free
dilutions is less animals
than 1 week
· stock
solutions
irritating to
skin and eyes
· stock must
be protected
from heat,
light
· gas phase:
respiratory
irritant, fire
hazard above
55?C
273 | P a g e
Table 3B - Summary of recommended concentrations, contact times, advantages and
disadvantages of non-halogen chemical germicides. The wide ranges of effective
concentrations and contact times cited reflect the interdependence of time and
concentration as well as factors such as resistance of the particular class or strain of
target microorganism(s) and desired effect. Also, some germicides are available in
combinations (e.g., glutaraldehyde/phenol or peracetic acid/alcohol mixtures) which are
synergistic whereby the components in combination produce a greater antimicrobial
effect than the sum of their individual effects.
Effective
Concentrations
Advantages Disadvantages
and Contact
Times
Examples of
Laboratory
Uses
Aldehydes:
Glutaraldehyde
· 0.5-2.5%
alkalinized
aqueous
solution
· broad
spectrum
· expensive
· pH,
temperature
dependent
· does not
corrode
· 2-30 mins; up metal
· pungent
to 12 hours to
· can
odour
kill all spores
tolerate
organic load · toxic: skin,
eye,
respiratory
tract irritant
· cold sterilant
and fixative
· surface
decontamination
· instruments,
equipment,
glassware
· activated
solutions have
less than 2week shelf life
Formalin (37%
aqueous
formaldehyde)
· 3-27%
formalin (110%
formaldehyde)
· broad
spectrum
· pungent
odour
· cold sterilant
and fixative
274 | P a g e
in 70-90%
alcohol
· 10-30
minutes
Formaldehyde
(gas)
Ethylene Oxide
Gas
· 1-3 hours
· 50-1200
mg/L
· 1-12 hours
·
· skin, eye and
inexpensive respiratory
tract irritant
· does not
corrode
· potential
metal
carcinogen
(animal
· can
studies)
tolerate
organic load · may require
24 hrs or more
to kill all
spores
· surface
decontamination
· as for
formalin
· as for
formulin
· effective
penetration
· flammable
· on site
decontamination
of biological
safety cabinet
HEPA filters
· broad
spectrum
· porr
penetration of
covered
surfaces
· instruments
and equipment
· enclosed areas
· flammable,
reactive
· heat or
moisture
sensitive
· no heat or · toxic:
supplies,
moisture
potential
instruments and
evolved
carcinogen and equipment
mutagen
· penetrates
packaging
· some
materials
sterilized items
may need more
than 24 hours
for outgassing
275 | P a g e
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