BIOLOGY 204/205 Advanced Genetics Laboratory
TABLE OF CONTENTS
Introduction…………………………………………………………………………….
MODULE 1: Recombinant DNA………………..…………………………………….
MODULE 2: Gene Expression………………………………..……………………….
MODULE 3: Screening a Plasmid Library Through Protein Expression…………
MODULE 4: Detection of the ALU Insertion; DNA Fingerprinting………………
MODULE 5: Microarray……………………………………………………………….
Appendix A: Solutions Guide……………..………………………………………….
Appendix B: Sterile Technique………………………………………………………..
Appendix C: Spread Plate Technique………………………………………………...
Appendix D: Pipet Use……………………..………………………………………......
Appendix E: Pipet Exercises………………………….……………………………......
Appendix F: PCR Reagents and Conditions……………………………...………….
Appendix G: Pierce Protein Assay……………………….…………………………...
Appendix H: DNA/Protein Markers……………………………………………...…..
Appendix I: GST Plasmid Map……………………………...…………………….......
Appendix J: Streak Plate Method……………………………...……………….……..
Appendix K: SDS-PAGE Assembly Instructions…………………….…….………...
Appendix L: Western Blot Assembly Instructions…………………………………..
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BIOLOGY 204/5
Advanced Genetics Laboratory I and II
--- Introduction --Module 1 Recombinant DNA/Bacterial Transformation
This module gives you some of the experience you would receive if you were to subclone a gene as a part of your research. That is, once you transform a bacterial line with
the plasmid that you isolate, you will need to demonstrate that you have made the
transfer of the correct gene.
Goals:
1. To purify a plasmid and transform E. coli with the plasmid.
2. To demonstrate that the transformants carry the plasmid by characterizing the
transformants’ phenotypes.
3. Analyzing the size of the DNA plasmid in a cracking gel.
4. Hybridization with the original plasmid in a Southern blot.
5. Amplify the gene inserted into the plasmid by PCR.
6. Sequence part of the plasmid.
Module 2 Gene Expression
This module allows you to determine whether a cell is expressing a gene of interest,
either GFP or GST. You will run the bacterial lysate on gels, stain with Coomassie blue
to look for a protein of the correct size and perform a Western blot to determine
whether the protein of interest was expressed.
Goals:
1. Confirm, using PCR, that the plasmid from the transformed bacteria (from
module 1) has the GST gene.
2. Show that bacteria with the gene for GST in the expression plasmid are
expressing GST using gel electrophoresis of the bacterial lysate.
3. Show that GST is expressed by the bacteria using a Western blot of the lysate.
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Module 3 Screening a Plasmid Library Through Protein Expression
This module lets you search for a gene of interest and demonstrate to us that you
retrieved the right gene –not unlike the process that you would go through in the lab.
Goals:
1. An antibody will be used to screen a human cDNA library for a clone that
expresses the particular gene product (in this case, GST).
2. Positive clones will be isolated, plasmid will be isolated and restriction digests
will be done to determine the insert size.
Module 4 Human Genetics: Detection of the ALU Insertion; DNA
Fingerprinting
This module allows you to carry out processes and procedures that a forensic scientist
might carry out.
Goals:
1. Isolation of your cheek cell DNA for polymerase chain reaction amplification.
2. Analysis of PCR products by agarose gel electrophoresis for the Alu insertions in
chromosome 8 and a polymorphism called a variable number tandem repeats
(VNTR) D1S80 from the noncoding region of chromosome 1.
3. Use the techniques you learned to help solve a “crime” by comparing DNA
“found” as evidence, and the DNA of the “victim” and the “suspects.”
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Biology 204/205 Advanced Genetics Laboratory
Grading Policy
Biology 204 and Bio 205 are four credit courses. You will complete Modules 1 & 2 and a
grant writing exercise during the fall semester for Bio 204. For Bio 205 in the spring
semester, you will complete two modules (Modules 3, 4 if you have already completed
1 and 2) and Module 5.
The format of the course is a short introduction and two formal meeting times per
week. The emphasis of the course is on experimental design, techniques, data gathering
and analysis. Work at the bench is given priority over work in a lecture setting. The
modules are designed to approach real situations in ongoing research projects.
Therefore, the modules are not necessarily designed to be finished in three hours. A
few labs will run long, taking 4-5 hours to finish. A few labs will be relatively short.
Students will sometimes need to return on an alternate day, usually at their own
convenience, to perform a short manipulation. Sometimes an experiment does not
work and it has to be repeated. Coming to class well prepared and following directions
carefully will cut down on potential mistakes!
Grading:
Your grade will be based on the following components, each with approximately equal
weight:
1. Performance in laboratory
2. Discussion of experiments in class
3. Notebook (these will be checked weekly)
4. Laboratory report
5. Grant proposal (Bio 204)
6. Final oral exam covering both lecture and laboratory material
Supplies:
You will need the following:
 Laboratory manual (available through the Biology Department)
 Bound laboratory notebook
 UV safety glasses
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Sharpie permanent marker
Transparent tape
Note: It is important to read over the procedures in the laboratory manual and the
corresponding information in the Molecular Biology Laboratory Atlas before coming to
class. Be prepared to start work after an introduction by the instructor or TA. Check
the laboratory calendar so that you know when each module will be done.
Laboratory Notebook Guidelines
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Bound notebook; no loose-leaf
Record in blue or black ink
Number all pages
Date all entries
Name, course number and email address should be on front cover
Reserve 3 pages at the beginning for the table of contents; keep up to date
Mistakes should be crossed out with a single line through the entry then initialed
Do not skip pages, do not rip pages out
Unused portions of a page should have a diagonal line drawn through the blank
portion
Each experiment should begin on a new page
All data, calculations and graphs should be entered directly into the notebook
Neat, orderly, complete
Your notebook should provide enough detail so that another Advanced Genetics
student could pick it up and repeat your procedure by following your entries. You
should include all of the following information: What was done and why, who
suggested it, who did it and when it was done, what results were obtained and what
conclusions were drawn.
NOTE: NOTEBOOKS WILL BE CHECKED WEEKLY AND GRADED
ACCORDINGLY!
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Laboratory Report Guidelines
You will be asked to write a formal report of one of the results from one of the lab
modules. You will prepare this report as you would prepare a manuscript for
publication, with introduction, methods, results, and discussion sections. To aid your
preparation of this report, you should go to the library early in the semester and find a
short article from Genetics from the last 5 years (download a pdf version or photocopy
from a paper journal, the library has both formats). The format in Genetics is
appropriate for your report. Below is a description of the content and length of each
section.
The report in its entirety should not exceed 10 pages in length. It should be printed
double-spaced, with no less than 1-inch margins. It must be in 12-point size in a
common font. Each section except the introduction should be started by its section
name, in bold type. At the head of the report, you should provide a title that indicates
which exercise you are writing about and your name. Whenever possible, you should
strive to write succinctly and in the active voice.
Abstract: 250 words summarizing the experiment.
Introduction: The introduction provides an overview of what the report is about,
including why the exercise was done (the goal of the exercise) and an explicit statement
of the hypothesis or hypotheses being tested. Background information about the
biology underlying the exercise should be included in the introduction. Recommended
length: 1.5 pages.
Methods: The methods section must be detailed enough to allow the reader to repeat
the exercise. You do not need to repeat the detailed description of the protocols in the
laboratory manual, but you should refer to the methods in the manual (Format:
Laboratory manual Page x-y) at the appropriate points. Recommended length: 3 pages.
Results: The results section reports upon what happened during the exercise. You
must include photocopies of the final gels and provide in tabular form other
measurements and data you collected. Each figure should have a brief descriptive
caption, and each table should have a title. However, it is not sufficient to simply insert
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these figures and tables. You must interpret your results in the text of the section, with
references to the appropriate figure or table (Format: Fig. 1, Table 2A). Recommended
length: 2 pages.
Discussion: In the discussion, you should briefly re-introduce the main goal or
hypothesis presented in the introduction, and then describe how your results are
related to the goal or hypothesis. In subsequent paragraphs, you should discuss any
failures to obtain results, and describe what you believe happened and what you would
do differently to correct each problem. This is your opportunity to show how well you
understand the molecular processes underlying the protocols! Recommended length: 2
pages.
Grant Writing Guidelines
The grant proposal must be based upon a novel concept that could be explored
within the technological and financial limitations of our laboratory, using model
organisms only. These limitations will aid you in narrowing your choice of topic.
A. Topic: The topic chosen should be novel, which means that no one else has worked on
this same exact problem before. However it should also be one that can realistically
attacked in the context of this laboratory. You should be able to put this topic into a
broader context; Why is this an interesting problem? What has already been done with
this problem in the past?
1. From this topic you must develop a testable hypothesis. This means you can
develop an experiment that will result in data that leads you to clearly be able to
reject or accept the hypothesis.
2. The experiment(s) will utilize techniques that you can do within this laboratory or
with our collaborators.
3. If you are currently working in a laboratory outside of this class the topic you
choose MUST be independent of that laboratory. You are not permitted to work on
the same model organism or a topic that is related to your outside research.
4. You are permitted, indeed encouraged, to discuss possible topics with classmates,
friends, family and other faculty.
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B. Model Organisms: Your experiments must be limited to classic model organisms from
molecular genetics that are readily available, easily maintained and for which we have
appropriate technologies to care for and manipulate. Examples include: Drosophila,
bacteria, Paramecium, C. elegans, yeast, small plants like grasses. You cannot work with
organisms that are difficult and expensive to maintain (i.e. mice or other small rodents,
fish, large vertebrates, humans, etc.)
C. Grant Format: Below is the format that your grant must follow. All text must be double
spaced 12 point type with 1 inch margins. Make sure your grant contains all of the
information within the guidelines given:
1. Cover Page - Fill out the cover page provided completely
2. Table of Contents - Page two is a table of contents. Provide the page number of
each category. Number pages consecutively at the bottom right of each page
throughout the application (including the cover page as page one).
3. Biographical sketch - A one page biographical sketch of the Principle Investigator
(PI; this is you!). This contains your name, your date of birth, your education
history, your previous relevant employment and a brief description of your prior
experience that allows the reviewer to recognize you are capable of doing the
proposed research.
4. Research Plan - This section should be 10 pages total. It should contain sufficient
information needed to evaluate the project, independent of any other
documentation. Be specific and informative, avoid redundancies. All tables, graphs,
figures, diagrams and charts must be included within the 10 page limit. The
following questions must be addressed in your research plan:
a. What topic will you be trying to address?
b. Why is this research important?
c. What has already been done in this field/topic?
d. What exact experiments will you be performing?
e. What are your expected results? (This is to clearly show your hypothesis.
What results will allow you to accept your hypothesis? What results will
cause you to reject your hypothesis?)
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5. Literature Cited - All references cited in the proposal (of which there should be
many) must be listed alphabetically by first author. They must include all authors,
year of publication, complete title of article, journal name (no abbreviations or
websites), volume and page numbers.
6. Appendix - In an appendix not to exceed 4 pages you may provide detailed
descriptions of your protocols, with all appropriate references in your citations*.
Your descriptions of the protocols should be as detailed as those provided in the
laboratory manual.
*Protocols: If you are using protocols in the laboratory manual you may cite the manual
itself as a reference. If, however, you propose other protocols you find and cite the original
reference that describes the protocol.
Use the following format and page distribution for section 4 - Research Plan:
a. Specific Aims: Clearly state your hypothesis and then list the Specific Aims of your
research to test the hypothesis. Specific Aims should organize the experiments that
you will do to test the hypothesis. (See example below.) List the broad, long term
objectives (goals) of the research and then present the proposed project's
relationship to these goals. This section should not exceed one page.
b. Background Information: Summarize the major research that has been done on this
topic leading to your proposal. Critically evaluate existing knowledge, and
specifically identify the gaps. State where your research will fit in with what has
been done previously, what new information does your research hope to discover?
This section should not exceed two pages.
c. Research Design and Methods: Describe the overall research design and the
specific procedures to be used. Include how data will be collected, analyzed and
interpreted. Discuss the potential difficulties and limitations of the proposed
procedures and alternative approaches to achieve the aims. Also provide a timeline
for the project (not to exceed six weeks.) This section should not exceed six pages.
d. Possible Results: In terms of the hypothesis you are proposing to test, state what
the results are expected. Which possible results will cause you to accept your
hypothesis? Which possible results will cause you to reject your hypothesis? What
are the limitations that may not allow you to have a clear answer? This section
should not exceed one page.
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This information was modified from the NIH instructions for US Department of Health
and Human Services Public Health Grant
http://grants.nih.gov/grants/funding/phs398/phs398.html
Example: Hypothesis is that the moon is made of blue cheese.
Specific Aim 1: Build a rocket ship to get to the moon and tools to sample cheese.
Specific Aim 2: Sample cheese on the moon and analyze it for its cheese-ness. Is it Blue or
Velveeta?
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Safety in the Laboratory
General Rules:
1. Disinfect your bench top with a 10% bleach solution when you arrive and when
you finish lab.
2. Wear gloves (lab coats, closed toe shoes and safety goggles are also highly
recommended). Avoid touching face and/or hair during an experiment.
Chemicals on your gloves can be transferred to your face/hair.
3. Wash your hands before you leave lab.
4. Do not eat, drink, smoke, chew gum or apply cosmetics while in lab.
5. Dispose of all used materials as directed.
6. Keep aisles clear.
7. Wipe all spills immediately. Inform lab tech and/or TA if you spill ANYTHING.
8. Dispose broken glass in the appropriate receptacle. Inform lab tech and/or TA
that you have broken glass.
9. Tie back long hair.
10. Try to avoid wearing baggy, loose clothing that can interfere with your
experiment and may catch on fire.
11. Extinguish burners as soon as you finish using them.
12. All Chemical Safety and MSDS information is located in the binder on the back
of the door.
13. If you are unsure about a procedure, just ask.
General Instructions for Recombinant DNA Work
The recombinant DNA involved in Modules 1 and 2 requires a P1 level of containment.
That means you must observe the following precautions:
1. Gloves must be worn to do the lab work.
2. Hands must be washed before leaving the laboratory
3. No eating, drinking or smoking in the laboratory.
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Pipetting :
1. Acquaint yourself with the various denominations of pipettors in an attempt to
avoid mistakes, particularly when working under time pressure.
2. The height of the fluid in the glass pipettes is measured at the bottom of the
meniscus while the pipette is being held vertically.
3. Never put a pipette back into a sterile container.
4. Do not handle the lower part of the pipette.
Serial Dilutions: Serial dilutions allow you to dilute a sample many fold by making a
series of small dilutions.
Standard Dilution Steps: Unless special circumstances demand it, the following are
the only dilution steps that are used (For convenience and error avoidance in
performing the accompanying arithmetic): 10, 20, 50 and 100.
10X
1:10
0.1 ml/0.9 ml
100 µL/900 µL
20X
1:20
0.1 ml/1.9 ml
50 µL/950 µL
50X
1:50
0.1 ml/4.9 ml
20 µL/980 µL
100X
1:100
0.1 ml/9.9 ml
10 µL/99 0µL
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Module 1
Recombinant DNA
Please refer to page 3 for introduction
**Note: The E.coli cell line used in this module is K12. The K12 bacteria cells are to be
transformed with the GST plasmid.
1.0 Overnight (ON )Bacterial Culture (Done for you)
1. The lab tech will add 2.5 ml of cells previously grown ON to 125 ml LB amp
medium (per group).
2. The cells will grow with shaking at 37C ON.
1.1 Isolation of Plasmid DNA
HAZARDOUS CHEMICAL INFO:
-Salt-Saturated Phenol is to be used only wearing gloves under the hood. Dispose of all
pipets and liquid waste containing SS Phenol in appropriate disposals.
-Chloroform: Isoamyl Alcohol (24:1) is to be used only wearing gloves under the hood.
Dispose of all pipets and liquid waste containing C:IA in appropriate disposals.
1. Transfer 125ml of the overnight bacterial culture (Transformed E. coli) to a large,
sterile centrifuge bottle and harvest the bacteria by centrifuging at 5000 rpm, 4ºC
for 10 minutes in the Beckman J2-21.
2. Decant the supernatant broth into the waste jar.
3. Resuspend the bacterial pellet in 5ml of Solution I containing 5mg/ml lysozyme.
4. Transfer to a 50ml polycarbonate Oakridge centrifuge tube. Let stand at room
temperature for 5 minutes.
5. Add 10ml of freshly made Solution II. Place the cap on the tube and mix the
contents by inverting the tube several times. Let stand on ice for 10 minutes.
6. While tubes are on ice, put RNase A in 100°C heat block for 15 minutes. Allow to
cool to room temperature before using in Step 11.
7. Add 8ml of ice-cold 5M potassium acetate (pH 4.8). Fill tubes only ¾ full. Screw
on the cap and mix by inverting. Let stand on ice for 10 minutes.
8. Balance the tubes before centrifugation.
9. Centrifuge in the Beckman J2-21; 15,000 rpm, 4ºC for 20 minutes. The genomic
DNA and bacterial debris should form a tight pellet at the bottom of the tube.
10. Being very careful not to disturb the pellet, divide the supernatant in half.
Transfer each half to separate 30ml glass tubes.
11. Add 500µL of heat treated RNase A to each tube.
12. Incubate at 37ºC for 20 minutes.
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13. In the chemical flow hood, add one volume of SS (salt saturated) phenol. (Note
the yellow color which helps you identify the phenol phase in the next step.)
Your tubes can be no more than 2/3 full including the addition of the phenol, so
divide your original solution as necessary into 3 or 4 conical tubes.
SAFETY NOTE: Phenol can cause severe burns to skin and damage clothing.
Gloves, safety glasses, and a lab coat should be worn when working with
phenol. All manipulations should be carried out in a fume hood. A glass
receptacle is available exclusively for disposing of used phenol and
chloroform.
14. Vortex the conical tube and contents with lids on for 1 minute; be sure the
contents are thoroughly mixed. Make sure the tops of the conical tubes are
screwed on tightly to ensure that no leaking will occur. Centrifuge for 1 minute
at 2800 rpm using the IEC Centra 7 Benchtop.
15. Transfer the non-colored upper, aqueous phase to a fresh conical tube. Do not
take the interface which is denatured protein. In the hood, add 1 volume of
chloroform: isoamyl alcohol (24:1). Vortex 1 minute and centrifuge 1 minute at
2800 rpm.
16. Transfer the upper, aqueous layer to a fresh 30ml glass tube and add 2.5 volume
of cold 95% ethanol. You need to calculate how much total liquid will be in each
30ml glass tube. The tube cannot be more than 2/3 full, so you may have to use
more than one 30ml glass tube. Make your calculations before adding the
ethanol!
17. Mix and allow to precipitate on dry ice for 15 minutes.
18. Balance your tubes along with their rubber sleeves.
19. Recover the DNA by centrifuging the tube at 4ºC in the Beckman J2-21 at 9500
rpm for 30 minutes.
20. Discard the supernatant into a waste container. The pellet will look like a whitish
residue on the side of the tube. To resuspend the pellet, use the pipette tip to
scrape the sides and then wash the residual ethanol up onto the sides of the tube
using a Vortexer. Try to resuspend the entire pellet to increase your plasmid
yield.
21. Start with one tube: wash the pellet with 1ml 70% ethanol by pipetting up and
down to resuspend the pellet. Transfer the solution from the first tube to the
resuspended pellet in another Oakridge tube and mix well. Transfer to the next
tube until all pellets are resuspended and pooled together. Transfer the solution
into one sterile 1.5ml microfuge tube.
22. Microcentrifuge for 5 minutes at 14,000 rpm. Discard the ethanol; add 1ml more
of ethanol to wash the pellet. Spin at 14,000 rpm for 5 minutes.
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23. Discard the ethanol; dry the pellet using the SpeedVac in the basement. Give
your sample to the TA/Lab tech to be properly dried.
24. Dissolve the pellet in 0.3ml TE. Aliquot 100 L to each of 3 microcentrifuge tubes
(properly labeled!).
25. Store at -20ºC.
1.2 Agarose gel to confirm isolation of the plasmid
HAZARDOUS CHEMICAL INFO:
-Ethidium Bromide is an extremely toxic carcinogen. WEAR GLOVES when handling,
and dispose of everything that has contacted EtBr in the appropriate solid waste
container.
-UV light is very harmful if looked at directly. When viewing your gels on the UV light
box be sure to wear a protective face mask, or place the shield on top of the box before
turning on light.
1.
2.
3.
4.
Prepare 250ml 1X TAE from 5X TAE stock.
Dissolve 0.35g agarose in 50ml 1X TAE buffer to make a 0.7% gel.
Microwave on high for 1 minute.
When done, swirl the flask and make sure all of the agarose is dissolved. If not,
microwave until it is.
5. Place the running tray into the gel-casting tray. Add comb.
6. Cool agarose slightly, approximately 5 minutes; slowly pour into gel casting set
up (from the farthest corner away from the comb) with comb and let cool until
opaque. Try to avoid bubbles!
7. While your gel is setting, thaw out one tube of your plasmid DNA on ice.
Just before you are ready to load the gel, heat the λ Hind III marker for 7
minutes in the 65ºC hot block.
8. Mix 4µL of 6X DNA sample buffer with 20µL plasmid DNA on a piece of
Parafilm.
9. Once your gel is set, remove it from the casting tray. Place it in the running tray,
with the comb still set. Cover the gel with 1X TAE. Gently remove the comb.
Removing the comb last will ensure that your wells do not collapse.
10. Be prepared to load the get quickly—you do not want your DNA to diffuse into
the running buffer.
11. Load 24µL of plasmid DNA sample and 20µL of λ Hind III marker in the wells;
put the lid on the box so that the DNA will run toward the red electrode.
12. Run the gel at 100V for ~1 hour.
13. Stain the gel for approximately 7 minutes in ethidium bromide, and destain in
water for 4 to 5 minutes.
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14. Examine the gel on the UV light box. If the ladder is not visible or is faint, place
the gel back into the stain. When you feel that your gel is properly stained, take
a picture to document your results. (Make a photo copy for your lab partner.)
15. Leave the gel in destain or discard (your TA will inform you).
1.3 Grow an overnight broth culture of E. coli (Done for you)
1.4 Transformation
Three hours before class the tech will take 1 ml of an ON culture and inoculate 50 ml
of fresh LB broth with it. It will shake at 37C for three hours. This will produce
exponentially growing cells for you to transform.
1. Divide broth culture into 2 sterile 50ml Oakridge centrifuge tubes; place tubes in
ice for 30 minutes.
2. Thaw out one tube of your plasmid DNA on ice.
3. Centrifuge the cultures at 4ºC in the Beckman J2-21 for 7 minutes at 5000 rpm;
decant the supernatant into the collection flask provided.
4. Resuspend one pellet in 25 ml ice cold 50 mM CaCl2. Combine this
resuspension solution with the second bacterial pellet; place on ice for 20
minutes. Keep CaCl2 on ice while waiting.
5. Centrifuge the cell suspension at 4ºC in the Beckman J2-21 for 7 minutes at 5000
rpm.
6. Decant the supernatant and resuspend the pellet in 3ml ice-cold 50 mM CaCl2;
place on ice for 5 minutes.
7. Dispense 2 aliquots of 0.3 ml cells in ice-cold microfuge tubes; add 0.2 ml of
transformation buffer to each tube.
8. Add 5 µL [>2 g] plasmid DNA to one tube. The second tube will not contain
plasmid DNA and will act as a control. Mix gently and leave on ice for 20
minutes.
9. Heat shock cells for 1 min in 42°C water bath.
10. Let stand at room temperature for 5 minutes.
11. Add 0.7 ml LB to each tube and tap gently with finger.
12. Shake at 37ºC for 60 minutes.
**NOTE: during this hour incubation your TA or Lab Tech will demonstrate
proper spreading and streaking procedures for plating. It is very important that
you understand sterile technique when working with bacteria so you don’t
contaminate your samples.
13. Plate 0.05, 0.1, and 0.3 ml of the cells with plasmid DNA onto LB amp plates.
Use the spread plate technique.
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14. Streak (Do not use the spread plate technique) the contents of the “no DNA
tube” on an LB amp plate and an LB plate. The LB amp plate will act as a
negative control, while the LB plate will serve as a positive control.
15. Label plates appropriately with group number, date, type of bacteria, and any
other important information, such as how much bacteria was plated.
16. Incubate the plates at 37ºC overnight (upside down); be sure to remove, wrap in
Parafilm and refrigerate the plates tomorrow!
Following Day: 1.5 Selecting for bacteria that carry the plasmid________
1. Examine transformed and no DNA control plates. (There should be no colonies
on the “No DNA” plate)
2. Choose 6 well isolated colonies from the transformed plates. Streak each colony
on half of an LB amp plate.
3. Choose 2 well isolated colonies from the control (non-transformed) plate
provided. Streak each colony on one half of an LB plate.
4. Incubate the plates overnight at 37ºC.
5. Wrap the old plates in Parafilm and refrigerate.
1.6_Secondary selection of transformed bacteria_____________________
1. Transfer 4 well-isolated colonies from 4 different transformed streaks and 2
control colonies into separate 1ml aliquots of sterile saline. Refrigerate the old
plates.
2. For the transformed bacteria, streak 1 loopful of saline/bacteria suspension onto
½ of an LB amp plate. Do this for each of the 4 samples.
3. For the control cells, streak 1 loopful of the saline/bacteria suspension onto ½ of
an LB plate. Be sure to label plates clearly!
4. Incubate at 37°C overnight; remove and refrigerate the next day.
5. Go to 1.16
1.7 Preparing bacteria for the cracking gel
(day before 1.8)
1. Using a marker, draw a line down the center of a new LB amp plate. Make a
template on paper with 1.5 cm x 1.5 cm squares on each half. Place the plate over
the template.
2. Using sterile tweezers, select a sterile toothpick.
3. Choose 2 LB amp plates from Day 1.6 that show the best growth. With the
toothpick, select one colony from the Day 1.6 plate and “fill in” the square on the
agar on the plate. Repeat for the 2nd colony using a new toothpick.
4. Repeat the procedure for the control, but use a fresh LB plate.
5. Incubate at 37ºC overnight for at least 24 hrs, but less than 36 hrs.
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1.8 Next day: Cracking gel
HAZARDOUS CHEMICAL INFO:
-Ethidium Bromide is an extremely toxic carcinogen. WEAR GLOVES when handling,
and dispose of everything that has contacted EtBr in the appropriate solid waste
container.
1. Make 250ml 1X TAE.
2. Prepare 0.7% agarose gel.
3. Use a sterile toothpick to scrape bacteria from the plates prepared the day before.
Add bacteria from each square to 250 µL of cracking buffer (Two squares for one
tube of 250 µL of cracking buffer). Do this for transformed and non-transformed
cells (you should have a total of 2 microcentrifuge tubes).
4. Incubate at 37ºC in the hot water bath for 25 minutes.
5. Centrifuge for 15 minutes at 14,000 rpm.
6. Use a toothpick to remove the bacterial debris from the bottom of each tube.
(You won’t be able to see a pellet, but when you pull it out, it will look like a blue
glob).
7. Load the gel:
8. Lane 1: 10 µL Hind III marker (Heat in 65C hot block for 7 minutes before
loading)
Lane 2: 10 µL plasmid DNA solution (5µL plasmid DNA + 2µL 6X DNA sample
buffer + 3µL 1X TAE)
Lane 3: Transformed supernatant
Lane 4: Non-transformed supernatant
Lane 5: 50µL Cracking buffer only
Note: Load as much transformed and nontransformed supernatant as possible
(A well formed well can hold ~50 L).
9. Run the gel for 1 hour at 100 volts.
10. Stain with ethidium bromide, destain, and photograph. Look for genomic DNA,
plasmid DNA and RNA.
1.9 Labeling DNA with Biotin
Part A: Labeling Reaction
1. Remove an aliquot of Plasmid DNA from the refrigerator and place on ice.
2. Add labeling reaction components to a 0.5ml tube (on ice) in the following order:
1X DNase I Buffer
19.9µL
DNase I Enzyme
0.1µL
dNTP mix
28µL
18
3.
4.
5.
6.
7.
Plasmid DNA
4µL
dH2O
128µL
DNA Polymerase I
20µL
Mix well and centrifuge for 5 seconds at 14,000 rpm.
Allocate 50µL into 4 tubes.
Incubate at 15°C for 2 hours in thermocycler.
Add 5µL Stop Buffer to each tube and mix.
Incubate tubes at 65°C for 5 minutes in thermocycler.
Part B: Purification of DNA probes
1. Transfer liquid to consolidate solution from 4 tubes into one tube.
2. Add 4µL 10% SDS to tube and mix.
3. Add 110µL Chloroform and 110µL SS Phenol to an empty 1.5ml
microcentrifuge tube.
4. Transfer DNA solution to chloroform phenol tube. Vortex 2 minutes and
then centrifuge for 2 minutes at 14,000 rpm
5. Collect the top layer of liquid and transfer to a fresh 1.5ml tube. Discard
remaining liquid into waste container.
6. Add 220µL chloroform to tube. Vortex 2 minutes and then centrifuge for 2
minutes at 14,000 rpm.
7. Repeat step 5.
8. Add 40µL 3M Sodium Acetate (pH 4.8) and 800µL cold 95% ethanol. Mix
gently by inverting tube.
9. Store at -20°C ON (at least 6 hours)
The Next Day:
10. Centrifuge for 5minutes at 14,000 rpm.
11. Carefully remove the supernatant.
12. Resuspend the pellet in 1ml cold 70% ethanol. Centrifuge for 5 minutes at
14,000 rpm.
13. Remove supernatant (ethanol). Let tube dry in cabinet for at least 1 hour.
14. Once dry, resuspend probe in 12µL TE buffer and store at -20° C.
1.10 Preparing for the Southern Blot (day before 1.11)
1. Using a marker, draw a line down the center of the underside of a fresh LB amp
plate. Draw two 1.5 cm x 1.5 cm squares on the underside of the plate, one on
each half.
19
2. Using a sterile toothpick, pick one isolated colony from the Day 1.6 LB amp
transformed plate. “Fill in” one square on the fresh LB amp plate with one
colony. Repeat for the second square making sure to use a fresh toothpick.
3. Repeat steps one and two, this time using an LB plate and the Day 1.6 nontransformed cells.
4. Incubate both plates for at least 24 hours.
1.11 Southern Blot
HAZARDOUS CHEMICAL INFO:
-Ethidium Bromide is an extremely toxic carcinogen. WEAR GLOVES when handling,
and dispose of everything that has contacted EtBr in the appropriate solid waste
container.
1.
2.
3.
4.
Run cracking gel (same as Day 1.8). Do not forget control lane!
Stain with ethidium bromide, briefly destain, and examine the gel.
Make sure to destain the gel for approximately 5 minutes before denaturing.
DO NOT cut the gel before running the southern blot. Nick a corner of the gel
before destaining for orientation purposes. Photograph the gel before destaining
completely—you will use this photograph later to compare to the results of your
southern blot.
5. Denature gel in 0.5 M NaOH/0.8 M NaCl for 30 minutes, rocking. Decant the
solution and repeat.
6. Rinse gel in dH2O for 1 minute.
7. While the gel is rinsing, cut and hydrate the nitrocellulose filter for 3 minutes in
dH2O, then in 10X SSC until blot set-up is ready. Make sure to notch the corner
of the nitrocellulose for orientation purposes and always wear gloves when
handling the nitrocellulose. Always handle the filter with forceps, and only
around the edges so as to not create blotches of background color.
8. Neutralize gel in 0.5 M Tris/1.5 M NaCl (pH 7.0) for 30 minutes, rocking. Decant
the solution and repeat.
9. Rinse the gel in 10X SSC for 3 minutes, rocking.
10. While the gel is neutralizing, prepare the Test Blot for 1.13.
a. Take your Biotin labeled probe out of the freezer and let thaw on
ice.
b. While thawing, cut a small piece of nitrocellulose and hydrate in
dH20 for 3 minutes.
c. Soak nitrocellulose in 10X SSC until the probe is thawed.
d. Using 2 µL of probe, make a small dot on the filter.
20
e. Dry in 80°C oven, then wrap in plastic wrap and store in the freezer
until 1.13.
11. Assembling the Southern Blot:
-First the wick (a long strip of paper towel will work) needs to be placed
on the platform so that it can only touch the buffer on two sides.
-Place three squares of Whatman 3M filter paper on top of wick.
-The gel should be placed on top of the filter paper, and the nitrocellulose
on top of that. **Make sure the nitrocellulose and the gel are lined up in the
correct orientation so you can compare them later**
-Place three more squares of Whatman 3M filter paper on top of the
nitrocellulose.
-A stack of cut paper towels at least 10 cm high should be assembled and
tied together with string. This can go on top of the filter paper.
-Pressure should be applied to the top of the stack to enhance wicking
overnight.
(Your TA should demonstrate this and assist in the assembly)
12. Let Southern Blot transfer ON in 10X SSC.
1.12 Drying of Blot (Done for you)_________________________________
1.
2.
3.
4.
Disassemble blot and rinse nitrocellulose filter in 5X SSC for 2 minutes.
Dry on large Kimwipe.
Bake filter in vacuum oven at 80ºC for 2 hours.
Wrap in plastic Wrap and store in freezer.
1.13 Hybridization of the Southern Blot
1. Denature 200L of Herring sperm DNA (2mg/mL) by boiling for 10 minutes
followed by fast chilling on ice.
2. While the Herring sperm DNA is boiling, soak the nitrocellulose filter (from
Days 1.11/1.12) and the test spot (from Day 1.11) in 2X SSC until uniformly
hydrated.
3. For prehybridization of the nitrocellulose filter, add the 200µL of freshly
denatured Herring sperm to the prehybridization solution. Mix, and then
transfer to two plastic tubs. Add the nitrocellulose filter to one, and the test spots
to another tub.
4. Incubate at 42ºC for 2 hours. The volume of prehybridization solution used
should be 20 to 100L per cm2 of the filter, ~7.5ml.
5. For hybridization, heat-denature the probe made on 1.11 and 200 L of Herring
sperm DNA by boiling for 10 minutes in a boiling water bath and then place on
ice. Just before use, add to the hybridization solution.
21
6. Remove the prehybridization solution from the tub and add the hybridization
solution to the filter (20-100 L per cm2), ~7.5 ml. The filter should be hybridized
at 42ºC overnight to achieve maximal sensitivity.
Following Day: 1.14 Detection of the DNA
**All the washes in this section need to be completed while rocking.**
Decant and save the hybridization solution in an appropriate size tube. Store at 4C.
1. Wash the filter & test spots with 100ml of 2X SSC/0.1% (w/v) SDS at room
temperature for 3 minutes. Decant the SSC and repeat.
2. Wash the filter & test spots with 100ml of 0.2X SSC/0.1 % (w/v) SDS at room
temperature for 3 minutes. Decant the SSC and repeat.
3. Wash the filter & test spots in 100ml of 0.16X SSC/0.1% (w/v) SDS at 50ºC for 15
minutes. Decant the SSC and repeat.
4. Rinse the filter & test spots in 100ml of 2X SSC at room temperature for 1 minute.
5. Dry on large Kimwipe and then wrap in plastic wrap and store in refrigerator.
6. The hybridization mixture containing the biotin-labeled probe may be reused.
Store the mixture at 4C for several days or at -20C for longer periods. Placing
the hybridization solution in a boiling water bath and cooling on ice just prior to
use should denature the probe.
1.15 Development of Blot
HAZARDOUS CHEMICAL INFO:
-NBT/BCIP is highly toxic. WEAR GLOVES when handling and dispose of all liquid
waste containing NBT/BCIP in the appropriate waste container.
1. Wash the filter and small test squares (from Day 1.11) in Buffer 1 at room
temperature for 1 minute with sufficient buffer to cover the blots. Decant Buffer
1 into the sink.
2. Incubate filters in Buffer 2 in a plastic container (1 hour, 65ºC) with sufficient
buffer to cover the blots.
3. Wash the filters in freshly made strep-avidin alkaline phosphatase (SA-AP)
conjugate for 15 minutes at room temperature.
(Add only enough SA-AP conjugate to cover the filters (~10ml). Use gentle agitation and
occasionally pipette SA-AP over the filters.)
4. Decant and save the SA-AP in a 15ml tube. Save for step #8.
Wash the filters in Buffer 1 using 20 to 40-fold greater volume than employed in
step 3. Gently agitate filter for 15 minutes in Buffer 1.
(ie., if you used 7 ml diluted SA-AP conjugate in step 3, wash with at least 140-280ml
Buffer 1.) Decant Buffer 1 into the sink.
5. Wash the filters for 10 minutes in Buffer 3. Decant Buffer 3 into sink.
22
6. Add 1ml NBT/BCIP solution to the saved SA-AP. A blue color should develop
overtime. Wear gloves when working with NBT-BCIP.
7. Decant Buffer 3 into the sink and add 9 ml of NBT-BCIP solution to the filters.
Allow the filters to develop for 15 minutes to 1 hour.
8. DNA bands will be most evident on only one side of the filter (check your notch
for correct orientation). Check your filter every 2 minutes to ensure that overdevelopment does not occur.
9. Once bands have developed, decant the NBT-BCIP solution in the appropriate
waste container and wash the filter in TE. This will terminate the color
development reaction. The TE can then be decanted into the sink.
10. To dry, bake at 80C in a vacuum oven for 1-2 hours.
11. Measure the photograph of the cracking gel, and compare the relative position of
the plasmid band to the results of the blot. Interpret your results.
12. Wrap filters in plastic wrap and label. The lab tech will photograph and
distribute the gels for your notebooks.
1.16 Designing Primers
5’-------------------------------------------------------------------------------------------------------3’
698 bp
GST
Partial DNA Sequence for GST (Read left to right, top to bottom):
5’……GTATTCATGTCCCCTATACTAGGTTATTGAAAATTAAGGGCCTTGT
310
GCAACCCACTCGACTTCTTTTGA……….…ATCCTCCAAAATCGGATCTGGT
960
TCCGCGTGGATCCCCGGGAATTCATCGTGACTGACTA………….……………..3’
The glutathione S-transferase protein consists of 232 amino acids. The sequence—using
the one-letter abbreviation for each amino acid—is shown below.
MSPILGYWKIKGLVQPTRLLLEYLEEKYEEHLYERDEGDKWRNKKFELGLEFPNLPYY
IDGDVKLTQSMAIIRYIADKHNMLGGCPKERAEISMLEGAVLDIRYGVSRIAYSKDFE
TLKVDFLSKLPEMLKMFEDRLCHKTYLNGDHVTHPDFMLYDALDVVLYMDPMCLD
AFPKLVCFKKRIEAIPQIDKYLKSSKYIAWPLQGWQATFGGGDHPPKSDLVPRGSPGI
HRD
23
Using this information, design the primers to amplify the GST gene. Once you have
designed the primers, fill out the oligonucleotide request form. The primers will then
be made on a DNA synthesizer.
1.17 PCR
1. Set up 7 - 0.5 ml PCR reaction tubes according to the PCR chart in Appendix F.
Read the chart carefully and make sure you add the correct amounts of reagents.
PCR is a very sensitive reaction and adding the incorrect amounts of reagents
may cause poor results. Appropriately label your tubes with your group
number and tube number!
2. Before mixing the reactants, you must calculate how much water must be added
to make a total of 50 L (including the Taq Polymerase). This is necessary
because the amount of plasmid that you add might differ from tube to tube.
3. If you add too much DNA, nonspecific amplification may occur—ask your TA
how much DNA to add based on the approximate concentration of your plasmid
samples.
4. Add all reactants, except the Taq, while the tubes are on ice.
5. Once all reactants (except Taq) are added to the tubes, spin them briefly to bring
all the liquid to the bottom of the tube. Note: Only spin tubes briefly (5 sec.), 0.2
ml tubes are thin-walled and can crack if microfuged for too long.
6. Lastly add the Taq polymerase.
7. Keep the tubes on ice until the entire class is ready to load the thermocycler.
8. The thermocycler will run for approximately 3hrs. After the 3hr. period is over,
the thermocycler will stay at a constant 4C until the tubes can be placed in the
refrigerator by the lab technician or TA. This will ensure that the PCR products
will not degrade.
1.18 Examining the PCR product
HAZARDOUS CHEMICAL INFO:
-Ethidium Bromide is an extremely toxic carcinogen. WEAR GLOVES when handling,
and dispose of everything that has contacted EtBr in the appropriate solid waste
container.
1. Make 300 ml 1X TAE.
2. Prepare a 2% agarose gel.
3. Remove 20µL of PCR product from each tube; add to 4 µL of sample buffer.
Store the remaining PCR product at 4ºC.
4. DO NOT HEAT 100 BP MARKER!!!
24
5. Load the PCR products onto gel along with 10 µL of 100 base pair ladder.
6. Run the gel for 1 hour at 100 volts.
7. Stain, destain, and photograph the gel.
1.19 Searches of the sequence using BLAST
(Basic Local Alignment Search Tool)
The plasmid DNA has been sequenced. You will receive a printout of the results. You
will analyze this information using a computer program called BLAST.
To access the program, go to http://www.ncbi.nlm.nih.gov/BLAST
25
Module 2
Gene Expression
Introduction
In the following series of experiments, you will not be using the transformed cells you
created in Module 1. K12 cells do not perform as well in expression experiments, so
BL21 cells will be used in Module 2. BL21 cells express the GST protein much clearer.
With these cells, you will induce the expression of the Glutathione-S-transferase (GST)
and run a SDS-PAGE (Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis) to
show that the protein was expressed. You will also perform a Western blot and use
antibodies to confirm the presence of the GST.
2.1: Preparation of bacteria for SDS-PAGE
(day before 2.2)
1. Streak an LB-amp plate with freshly transformed BL21+GST cells from your lab
tech.
2. Streak an LB plate with control (non-transformed) BL21 cells from your lab tech.
2.2: SDS-PAGE preparation and Pierce Protein Assay
HAZARDOUS CHEMICAL INFO:
-Acrylamide is a neurotoxin. WEAR GLOVES when handling, and dispose of all
solid waste (pipets, gloves, or anything else that has come in contact with the
acrylamide) in the appropriate waste container.
-TEMED is highly toxic. WEAR GLOVES when handling, and dispose of all solid
waste (pipets, gloves, or anything else that has come in contact with the TEMED) in the
appropriate waste container. Do not inhale fumes.
-Ammonium Persulfate is highly toxic upon contact with skin. WEAR GLOVES
when handling, and dispose of all solid waste (pipets, gloves, or anything else that has
come in contact with the TEMED) in the appropriate waste container.
-N-Butanol is flammable. Use caution.
Three hours before class …inoculate bacterial cultures:
1. Inoculate 2 x 2 ml of LB amp broth with transformed bacteria (1 colony each).
2. To ONE of the LB amp cultures, add 60 L of 100mM IPTG.
3. Inoculate 2 ml of LB broth with control (non-transformed) bacteria.
4. Shake at 37C for 3 hours.
At lab time:
5. Obtain a large and small glass plate; wipe with methanol and a Kimwipe until
you hear a “squeaky” noise. Handle glass plates at edges. Wear gloves!
6. Assemble the gel casting apparatus (See diagram in appendix).
26
7. Insert the comb and use a Sharpie to draw a line across the glass 1cm below the
comb. Once the line is drawn, remove the comb.
8. Test to see if the apparatus is leak-proof. Squirt some water in between the glass
plates and look for leaks. If leaks occur, a tighter seal must be achieved. Placing
several layers of paper towels on top of the rubber seal can do this. Make sure to
remove the water before pouring the gel.
Before you pour your gel, have the TA or lab tech check your apparatus!
9. Prepare the resolving gel in a 15 ml tube according to the following directions.
DO NOT ADD THE FRESHLY MADE AMMONIUM PERSULFATE UNTIL YOU
ARE READY TO POUR THE GEL
Sterile dH2O
3.29 ml
4X resolving Buffer pH 8.9
2.60 ml
30% Acrylamide stock
4.00 ml
TEMED
10 L
Last:
Fresh 10% Ammonium persulfate 100 L
10. Gently swirl the solutions to mix WELL.
11. Using a Pasteur pipette, pour the gel by allowing the acrylamide solution to run
down along the side of the spacer. Add the acrylamide solution until it is just
barely above your Sharpie line. Try to avoid making bubbles.
12. Overlay the acrylamide with N-Butanol to aid in the polymerization. Do this by
gently adding the N-Butanol with a Pasteur pipette. You will be able to see a
distinct line between the N-Butanol and the 4X Resolving solution.
13. Allow the gel to polymerize for AT LEAST 30 minutes (Any extra acrylamide
mix in your tube will be a good gauge for polymerization. Make sure the cap is
on).
14. While your gel is polymerizing, you can prepare your samples and standards for
the Pierce Protein Assay if you have not done so already. Go to next section for
instructions.
15. Once your gel has polymerized, pour off the N-Butanol into the sink and rinse
with dH2O. Add a layer of dH2O on top of your gel using a Pasteur pipette.
16. Wrap the gel/casting apparatus in a damp paper towel and then plastic wrap.
Label appropriately. Store in the cold room.
Prepare the proteins for the gel and protein assay:
1. Split each culture into two 1.5 ml. You will have a total of 6 tubes (2 x
Transformed+IPTG, 2 x Transformed-IPTG, 2 x Untransformed).
2. Spin all tubes in the microcentrifuge for 1 minute.
3. Decant the supernatant from each tube.
27
4. Resuspend one of each kind of pellet (i.e. transformed, transformed plus IPTG
and control) in 50 L lysis buffer.
5. Transfer this solution to its complementary tube. Do this for each type of pellet.
You will now have a total of 3 tubes.
6. Sonicate each sample in ice, 3 times at 10-second intervals. Sonicating breaks
open the cells by sound waves.
EAR PROTECTION REQUIRED!
When sonicating, avoid touching the sides of the tube with the tip of the
sonicator. The sample may become frothy; try to keep the sample from coming
out of the tube. Turn off the sonicator and rinse the tip with dH2O in between
samples and wipe with a Kimwipe.
7. Record the approximate total volume of each sample.
8. Take 10 L samples from each sonicate; keep on ice for protein assay.
9. Freeze the remaining samples at -20C. Label appropriately!
Pierce Protein Assay (See directions in Appendix G)
Use the chart provided to develop a standard curve using BSA standards and to
determine your own protein concentration. **NOTE: Do the entire PPA and standard
curve development in DUPLICATE. Make two of every sample (label 1a, 1b, 2a, 2b, etc)
and average the OD readings at the end for a more accurate standard curve.
1. Dilute protein sample: Your protein samples should be diluted 5:95. To do this,
add 5 L of your sample and 95 L of sterile dH2O.
2. Make dye solution: Use Solutions A and B from the Pierce Protein Assay Kit.
They should be mixed 50:1…but make up only the amount you will need (~40
ml). Mix the dye in a 50 ml tube.
3. Add BSA and dH20 according to the directions in Appendix G.
4. Add 2ml of the dye to each one of your samples and standards. Vortex.
5. Incubate at 37C for 30 minutes.
6. Get OD values for standards and samples: TA will assist in the operation of the
spec.
7. Place your standard into a clean cuvette.
To clean the cuvette, rinse with dH2O. Make sure to dry the outside of the
cuvette with a Kimwipe. Handle the cuvette only on the frosted sides.
8. Read OD at 562nm.
9. Repeat for each standard and sample, including duplicates.
If only using 1 cuvette, make sure to rinse with dH2O between each standard.
10. Once the standards are complete, read your samples. You should blank the
instrument with dH2O and dye. If using only 1 cuvette, make sure to rinse with
dH2O between each sample.
11. Find the average of each standard and sample duplicates.
28
12. Establish a standard curve using the OD values obtained with your BSA
standards: graph OD (x-axis) vs. concentration (y-axis) on graph paper. Using
this graph, calculate the protein concentrations in your three samples.
13. Use Excel to plot your data on a second chart (This is homework). Make sure to
paste your Excel chart in your notebook properly labeled.
2.3 SDS-PAGE
HAZARDOUS CHEMICAL INFO:
-Acrylamide is a neurotoxin. WEAR GLOVES when handling, and dispose of all
solid waste (pipets, gloves, or anything else that has come in contact with the
acrylamide) in the appropriate waste container.
-TEMED is highly toxic. WEAR GLOVES when handling, and dispose of all solid
waste (pipets, gloves, or anything else that has come in contact with the TEMED) in the
appropriate waste container. Do not inhale fumes.
-Ammonium Persulfate is highly toxic upon contact with skin. WEAR GLOVES
when handling, and dispose of all solid waste (pipets, gloves, or anything else that has
come in contact with the APS) in the appropriate waste container.
-Coomassie Blue Stain and Coomassie Blue Destain are highly flammable and
irritating to the skin. WEAR GLOVES when handling and dispose of in appropriate
waste container.
1. Remove the resolving gel from the cold room and pour a 4% stacking gel.
2. Mix the following components in a 15 ml tube.
3. Sterile dH2O
6.10 ml
4X stacking buffer pH 6.8
2.50 ml
30%
Acrylamide stock
1.30 ml
TEMED
10 L
Last:
Fresh 10% Ammonium persulfate 50 L
4. Before adding the ammonium persulfate, pour the dH2O off the resolving gel
and dry with a Kimwipe.
5. Add the ammonium persulfate to your tube. Mix gently.
6. Pour the stacking gel as you did the resolving gel all the way to the top of the
small glass plate. If it overflows when inserting the comb this is okay. Clean the
comb thoroughly with methanol before inserting.
7. Being careful to avoid making air bubbles, insert the clean comb until there is no
air between the wells. This is VERY IMPORTANT; the stacking gel will not
polymerize if the comb is not clean, or if there is air between the wells. Ask your
lab tech to double check your set up.
8. Allow 30 minutes for the gel to polymerize. Thaw your protein samples on ice
while waiting.
29
9. Once the gel is set, remove them from the casting stand and assemble in the gel
box.
Note:
Do not remove the comb yet.
10. Add ~115ml of PAGE Running buffer to the upper chamber. The buffer level
should be half way between the big and small glass plate.
11. Add enough PAGE Running buffer to the lower chamber to cover the bottom
half of the gels.
12. Carefully remove the comb.
Gel Set Up
1. Calculate the volumes of sample, sample buffer, and water needed for each tube
(make 60 µl total so that even if some evaporates during boiling, there will still
be 40 µl left):
-Each tube needs 60 µg of protein total. Using the concentrations you
calculated off of your standard curve, calculate how many microliters
equals 60 µg.
-Each tube needs 1X sample buffer. You are given 6X sample buffer.
Calculate how much 6X sample buffer is needed so that the final
concentration is 1X.
-Each tube needs a total volume of 60 µl. Figure out how much water
should be added to each so that the total volume is 60 µl.
2. Place the remaining protein samples in the freezer.
3. Boil samples for 5 minutes right before you are ready to load.
4. Load you samples into the gel in the following order (use gel loading tips):
Lane 1: 6X SDS sample buffer/Blank
Lanes 2, 6, 10: Prestained protein marker (20 L per lane)
Lanes 3, 7: Transformed
Lanes 4, 8: Transformed plus IPTG
Lanes 5, 9: Control (non- transformed)
5. Run the gel at 50mA for 1-1 ½ hours.
6. Remove gel carefully from the gel apparatus. Use a razor blade to cut the
stacking gel portion away. Dispose of the stacking gel in the appropriate waste
container.
7. Put the gel into a plastic container and cover with Coomassie stain. Microwave
on low for 1 minute. Discard stain in appropriate waste container.
8. Transfer gel to destain. Wash and discard destain in correct waste container.
9. Submerge the gel in more destain. Shake gently overnight.
2.4 Dry Gel: Done for you
1. Discard destain in appropriate waste container.
30
2. Rinse gel in dH2O water. Discard into appropriate waste container.
3. Submerge your gel in more dH2O and place back on the shaker.
4. Your TA/Lab tech will appropriately dry your gels using cellophane and give to
you during the next lab meeting.
5. Make sure to observe and record gel appropriately.
2.5 Pouring a Resolving Gel for SDS-PAGE and Western Blot
HAZARDOUS CHEMICAL INFO:
-Acrylamide is a neurotoxin. WEAR GLOVES when handling, and dispose of all
solid waste (pipets, gloves, or anything else that has come in contact with the
acrylamide) in the appropriate waste container.
-TEMED is highly toxic. WEAR GLOVES when handling, and dispose of all solid
waste (pipets, gloves, or anything else that has come in contact with the TEMED) in the
appropriate waste container. Do not inhale fumes.
-Ammonium Persulfate is highly toxic upon contact with skin. WEAR GLOVES
when handling, and dispose of all solid waste (pipets, gloves, or anything else that has
come in contact with the APS) in the appropriate waste container.
1. Pour a 12% polyacrylamide resolving gel:
2. Obtain a large and small glass plate; wipe with methanol and a Kimwipe until
you hear a “squeaky” noise. Handle glass plates at edges.
3. Insert the comb and use a Sharpie to draw a line across the glass 1cm below the
comb. Once the line is drawn, remove the comb.
4. Test to see if the apparatus is leak-proof. Squirt some water in between the glass
plates and look for leaks. If leaks occur, a tighter seal must be achieved. Placing
several layers of paper towels on top of the rubber seal can do this. Make sure to
remove the water before pouring the gel.
5. Before you pour your gel, have the TA or lab tech check your apparatus!
6. Prepare the resolving gel in a 15ml tube according to the following
directions…BE SURE TO WEAR GLOVES…ACRYLAMIDE IS A
NEUROTOXIN…DO NOT ADD THE AMMONIUM PERSULFATE UNTIL YOU
ARE READY TO POUR GEL
Sterile dH2O
3.29 ml
4X resolving Buffer pH 8.9
2.60 ml
30% Acrylamide stock
4.00 ml
TEMED
10 L
Last:
Fresh 10% Ammonium persulfate
100 L
7. Gently swirl the solutions to mix.
31
8. Using a Pasteur pipet, pour the gel by allowing the acrylamide solution to run
down along the side of the spacer. Add acrylamide until it is just barely above
your Sharpie line. Try to avoid making bubbles.
9. Cover the acrylamide gel solution with N-Butanol. To do this, use a Pastuer
pipet and gently place N-Butanol over the top of the gel. Once gel has
polymerized, decant the N-Butanol and add a layer of dH2O. Wrap the gel
apparatus in a damp paper towel and then Saran Wrap. Label appropriately.
Store in cold room.
2.6 SDS-PAGE gel and Western Blot
HAZARDOUS CHEMICAL INFO:
-Acrylamide is a neurotoxin. WEAR GLOVES when handling, and dispose of all
solid waste (pipets, gloves, or anything else that has come in contact with the
acrylamide) in the appropriate waste container.
-TEMED is highly toxic. WEAR GLOVES when handling, and dispose of all solid
waste (pipets, gloves, or anything else that has come in contact with the TEMED) in the
appropriate waste container. Do not inhale fumes.
-Ammonium Persulfate is highly toxic upon contact with skin. WEAR GLOVES
when handling, and dispose of all solid waste (pipets, gloves, or anything else that has
come in contact with the APS) in the appropriate waste container.
-Western Blot Transfer Buffer is flammable. Wear gloves and use caution when
handling.
1. Remove the resolving gel from the cold room and pour a 4% stacking gel.
2. Mix the following components in a 15ml tube.
Sterile dH2O
6.10 ml
4X
stacking buffer pH 6.8
2.50 ml
30%
Acrylamide stock
1.30 ml
TEMED
10 L
Last:
Fresh
10% Ammonium persulfate
50 L
3. Before adding the ammonium persulfate, remove the dH2O and dry with a
Kimwipe.
4. Add the ammonium persulfate. Mix gently.
5. Pour the stacking gel as you did the resolving gel all the way to the top of the
small glass plate. If it overflows when inserting the comb this is okay. Clean the
comb thoroughly with methanol before inserting.
6. Being careful to avoid making air bubbles, insert the clean comb until there is no
air between the wells. This is VERY IMPORTANT; the stacking gel will not
32
polymerize if the comb is not clean, or if there is air between the wells. Ask your
lab tech to double check your set up.
7. Allow at least 30 minutes for the gel to polymerize. Thaw protein samples on ice
while waiting.
8. Once the gel is set, remove them from the casting stand and assemble in the gel
box.
Note: Do
not remove the comb yet.
9. Add ~115 ml of PAGE running buffer to the upper chamber. The buffer level
should be half way between the big and small glass plate.
10. Add enough PAGE running buffer to the lower chamber to cover the bottom half
of the gels.
11. Carefully remove the comb.
Gel Set Up:
1. Calculate the volumes of sample, sample buffer, and water needed for each tube.
This should be the same as in 2.3.
2. Place the remaining protein samples in the freezer.
3. Add 6X SDS sample buffer to each protein sample.
4. Boil samples for 5 minutes right before you are ready to load.
5. Load you samples into the gel in the following order:
Lane 1: 6X SDS sample buffer/Blank
Lanes 2, 6, 10: Prestained protein marker (10 L per lane)
Lanes 3, 7: Transformed
Lanes 4, 8: Transformed plus IPTG
Lanes 5, 9: Control (non- transformed)
6. Run the gel at 50mA for 1-1 ½ hours and then set up Western blot.
Blotting Procedure
The transfer will be accomplished using the Hoeffer Semi Dry Transfer Apparatus. This
unit transfers proteins from a polyacrylamide gel to a nitrocellulose membrane by
means of a low current and low voltage transfer.
1. Rinse the anode and cathode of the transfer apparatus with dH2O. Be careful not
to get the leads or interlock housing wet.
2. Prepare the gel for transfer. Carefully cut away stacking gel with a razor blade.
Measure the gel and record the dimensions.
3. Cut a hole in a Mylar mask 2 mm smaller than the gel. Center the mask on the
anode of the transfer apparatus.
4. Cut six pieces of blotting paper and one piece of nitrocellulose membrane the
same size as the gel. Measure and cut carefully; they must not be larger than the
gel! Make sure to notch the nitrocellulose for orientation purposes!
33
5. Soak the blotting paper in Western blot transfer buffer.
6. Rinse the nitrocellulose membrane with dH2O, then soak it in Western blot
transfer buffer for 5 minutes.
7. Put one piece of the blotting paper over the opening in the Mylar mask. Roll a
test tube over the paper 3-4 times to push all air bubbles out. You will need to
use moderate pressure to be effective.
8. Repeat this process adding two more blotting paper layers.
9. Add the nitrocellulose paper to the stack and roll out the air bubbles.
10. Add the gel. Do not roll. Be careful setting the gel on the stack. Try to line it up
correctly the first time as some proteins may stick to the membrane on contact
and moving the gel around will affect the quality of your blot.
11. Add the last three strips of blotting paper, one at a time, carefully rolling out the
bubbles each time.
**NOTE: If it is too hard to place the gel exactly on top of the nitrocellulose
membrane, you can put the gel down first and then the nitrocellulose, but you need
to remember to FLIP THE STACK after you have finished putting the filter paper on
top so that the nitrocellulose is BELOW the gel.**
12. Put the top on the transfer unit. Set a flask with 1L of water on top to add
pressure.
13. Connect the short safety interlock lead on the cover to the jack on the base. Plug
the leads into the power supply.
14. Turn on the power (0.8 mA per cm2 of gel surface). Transfer for 1½ hour.
15. Turn off the power supply. Disconnect the leads and safety interlock.
16. Use forceps to remove the nitrocellulose membrane. Place it face up on fresh
blotting paper. Record the orientation of the lanes, and then wrap the
nitrocellulose in plastic wrap and store at -20C. Blotting papers and gel can be
thrown away.
2.7 Primary Antibody Blocking (day before 2.8)
1. Cut the nitrocellulose through the center lane of the prestained marker. Wrap
one half of the blot in Saran wrap, label, and place back into the freezer.
2. Submerge the other half of the blot with 25ml of blocking solution. Rock at room
temperature for 1 hour.
3. Decant the blocking solution into the sink.
4. Wash the blot with 50ml TBS. Repeat.
5. Add the primary antibody (Anti-GST produced in rabbits diluted 1:5000 in 10 ml
of solution containing TBS, 0.1% Tween-20, and 1% dry non-fat milk). Add
enough to submerge the nitrocellulose filter.
6. Rock in the cold room overnight.
34
2.8 Staining and Detection of Western Blot
HAZARDOUS CHEMICAL INFO:
-Amido Black Stain and Destain are flammable irritants. Wear gloves when
handling and dispose of in proper waste containers.
Staining with Amido Black:
1. Remove nitrocellulose blot from the freezer and cover with a minimal amount of
amido black.
2. Rock at room temperature for 5 minutes.
3. Decant stain back into its original container then wash the blot with amido black
destain until all background color is gone.
4. Decant the destain into the proper waste container.
5. Place the blot on filter paper to air dry.
6. Observe the stained blot. Save it to compare to the immunostained half. Wrap
the blot in plastic wrap. Copies of the stained blot should be made for each
group member. Scanning and printing the blot can achieve this.
Detection of the antibody:
1. Decant the primary antibody into the sink.
2. Briefly wash the blot with TBS-T. Decant the TBS-T into the sink. Repeat 3X.
3. Add the secondary antibody (Goat anti-rabbit IgG alkaline phosphatase
conjugated diluted 1:10000 in TBS-T). Save a small volume of the secondary
antibody in a 1.5 ml tube to use as a control, keep on ice.
4. Rock the blot at room temperature for 1 hour.
5. Decant the secondary antibody into the sink.
6. Wash the blot with 50 ml of TBS-T. Rock at room temperature for 5 minutes.
7. After 5 minutes, decant the TBS-T into the sink, add fresh TBS-T and repeat for a
total of (6) 5 minute TBS-T washings.
8. Add 500 L of the NBT-BCIP to the secondary antibody you saved. A blue color
should develop within a couple minutes.
9. Add 10 ml NBT-BCIP solution directly to your blot. Agitate until color develops.
10. Once color has developed, decant the NBT-BCIP solution into its appropriate
waste container. Rinse the nitrocellulose with dH2O. Decant into the sink.
11. Air dry the nitrocellulose on filter paper. Observe and record. Wrap the
nitrocellulose in plastic wrap and store in your notebook. Copies of the blot
should be made for each group member.
35
Module 3
Screening a Plasmid Library Through Protein Expression
Introduction
In order to clone genes, scientists often turn to huge libraries of plasmids that have a
variety of sequences inserted into them, one sequence per plasmid. The trick is to pick
out the one that carries your gene. To find the gene, we can use a plasmid that has a
promoter before the insert and that expresses the mRNA for the inserted genes. The
mRNA is translated into protein. In order to find the bacteria expressing you gene, you
can use antibodies against the protein gene product. This experiment is designed to
screen a library for the GST or GFP gene (your TA will inform you of the choice).
You will grow the library bacteria on agar and screen the clones for the ones expressing
GST. Expression by individual clones is detected by overlaying the plates with
nitrocellulose filters that have been soaked with isopropylthio--galactoside (IPTG) to
induce protein expression. The filter is then incubated with anti-GST antibodies and the
GST producing colonies identified. The GST producing clone is then isolated and a
pure culture grown. The plasmid is isolated and a restriction enzyme digest is done to
cut out the receptor gene insert. The products from the digest are run on an agarose gel
for analysis.
3.1 Plating the bacteria
_
1. Serially dilute an E. coli culture that is the library with sterile dH2O so that at
least one plate has between 100 and 200 colonies. Use four tubes with the
following amounts of dH2O: 100 L, 150 L, 200 L, 250 L. Ask your TA how
much library to add.
2. Plate 100 L of your diluted bacteria on four properly labeled LB amp plates.
3. Incubate plates upside down overnight at 37C.
3.2 Overlaying the Plates
1. Pre-soak two nitrocellulose membranes in 10 mM IPTG and air-dry before use.
Gently handle the NC Paper with forceps. Soak the membranes separately.
Dispose of the IPTG in the appropriate waste container.
2. Carefully overlay the plates with the nitrocellulose filter.
3. Incubate at 37C for 3 hours right side up.
4. Using a needle and India ink, mark the filters. Choose an asymmetric pattern so
you will be able to match the filter and the plate later. Make the mark at the edge
of the plate. Make sure the ink marks the plate as well as the filter.
36
5. Remove the nitrocellulose filter and soak in ~7 ml of TBST. Wrap in plastic wrap,
label and store in refrigerator/cold room. Store LB amp plates in refrigerator/cold
room.
3.3 Identifying positive colonies
HAZARDOUS WASTE INFO:
-NBT/BCIP is extremely toxic. WEAR GLOVES when handling and dispose of all
liquid waste containing NBT/BCIP in proper waste container.
1. Incubate the nitrocellulose filters in 10 ml of blocking buffer in Petri dishes. Rock
at room temperature for 1 hour. Wash filters in separate containers.
2. Decant the blocking buffer and wash the filters with 6-10 ml TBST.
3. Decant the TBST and add 10 ml of primary antibody (Anti-GST produced in
rabbits, diluted 1:5000 in a solution containing TBS, 0.1% Tween, 1% dry non-fat
milk). Rock at room temperature for 1 hour. When done, decant and save the
antibody in the refrigerator.
4. Wash the nitrocellulose filter for 5 minutes in 6-10 ml TBST; rock at room
temperature. Repeat for a total of three washes.
5. Incubate the nitrocellulose filters in 10 ml of secondary antibody (Goat antirabbit IgG diluted 1:10000 in TBS-T). Rock at room temperature for 45 minutes.
When finished, save the secondary antibody for a check below.
6. Wash the nitrocellulose filter for 5 minutes in 6-10 ml of TBST; rock at room
temperature. Repeat for a total of three washes.
7. Decant the TBST and add 6 ml of the NBT/BCIP solution to the filter. Add 500
L of NBT/BCIP to the secondary antibody you saved as a control to check for
color development (a blue color should develop).
8. Once color has developed on the nitrocellulose filter, decant the NBT/BCIP in the
appropriate waste container and soak in stop solution for 10 minutes. Air-dry
the filter. Wrap the filters in plastic wrap and store in the cold room.
Wednesday 3.4 Isolation of positive colonies
1. Note positive colonies on the NC and find the corresponding colonies on the
agar plate.
2. Use a Pasteur pipet (or sterile toothpick) to select four positive colonies and 1-2
negative colonies for control
37
3. Put the agar plug in a tube containing 10ml of LB amp broth. Incubate overnight
in a 37C shaking waterbath.
3.5 Mini prep plasmid isolation
Note: Appropriately label all tubes and matching columns so there is no confusion!
Part I. Preparing the lysate:
1. Transfer 1.5 ml of bacterial culture into 2 microfuge tubes; do this for each of
your tubes.
2. Centrifuge at 12,000 rpm for 3 minutes at 4C.
3. Decant as much as the supernatant as possible. Resuspend the pellet in 200 L
cell resuspension solution. Transfer this mix to the matching tube and resuspend
the second pellet.
4. Add 200 L cell lysis solution and invert 4 times to mix.
5. Add 200 L neutralization solution and invert 4 times to mix.
6. Centrifuge lysate for 5 minutes at 13,000 rpm.
Part II. Plasmid DNA Purification:
1. Obtain 1 mini-column and 1 syringe for each culture. Remove the plunger from
the syringe and attach the column to the syringe. Place the column in a small test
tube.
2. Shake the resin bottle to resuspend the resin then add 1 ml to each minicolumn/syringe assembly.
3. Carefully transfer the cleared lysate from #6 above to the resin.
4. Insert the plunger and push the resin/lysate into the minicolumn.
Part III. Washing:
1. Detach the mini-column from the syringe. Remove the plunger from the barrel.
Reattach the mini-column.
2. Add 2 ml column wash solution. Insert the plunger and push the column wash
through the mini-column.
3. Remove the syringe and transfer the mini-column to a 1.5 ml microcentrifuge
tube. Centrifuge at 13,000 rpm for 2 minutes.
Part IV. Elution:
1. Transfer the mini-column to a new microcentrifuge tube.
2. Add 50L sterile dH2O directly into the column and wait 1 minute.
3. Centrifuge at 13,000 rpm for 20 seconds.
4. Remove and discard mini-column. Store DNA at -20C. Label tubes
appropriately!
38
3.6________________________
1.
2.
3.
4.
Remove one DNA sample from freezer and thaw on ice.
Transfer 18 L of the DNA sample to a clean microcentrifuge tube.
Add 2 L of 10X React 3 Enzyme buffer to the tube.
Add 1 L of BamH1 and 1 µL of EcoRV to the tube. Mix with pipette man.
Enzymes need to be kept on ice. Enzymes should be added last to well mixed
solutions.
5. Incubate at 37C for 45 minutes.
6. Microfuge at 12,000 rpm for 5 seconds.
7. Add 5 L of sample buffer to the digests.
8. Remove 10 L of the corresponding non-digested plasmid DNA to a clean
microfuge tube. Add 4 L of sample buffer and 10L TE.
9. Load onto a 0.7% agarose gel:
Lane 1:  Hind III marker. Heat at 65C for 7 minutes before loading onto gel!
Lane 2: Non-digested plasmid
Lane 3: Digested plasmid
9. Run at 100 v for approximately 1 hour.
10. Stain the gel in ethidium bromide, destain and photograph. Record and interpret
results.
39
Module 4
Human Forensic DNA Analysis
Introduction
This module allows you to analyze some of your own DNA for markers that are used in
forensic analysis and also to simulate the analysis of criminal evidence. One marker
that you will use is the TPA-25 sequence from an intron of the tissue plasminogen
activator gene. This is a member of the Alu family of insertions in our genomes. You
will amplify regions of chromosome 8 to determine whether the TPA-25 insert is in the
intron. We can screen for this insertion sequence using polymerase chain reaction
(PCR) because it has no phenotypic consequence, i.e. no health or other problems arise
from its presence.
We will also amplify a region of chromosome 1 region D1S80 to look for a polymorphic
sequence, that is a region that has a variable number of repeats (VNTR). Again, we will
use PCR to amplify a region and compare results among lab members to see how we
can identify each one uniquely by the size (14-40 repeats) of the PCR product from this
region. The sequence we will amplify is in a non-coding region and has no phenotypic
consequences.
The DNA template for your PCR will be your own, from a swab of your mouth.
The following primers are used to bracket the TPA-25 locusThe forward primer:
5’-GTAAGAGTTCCGTAACAGGACAGCT-3’
The reverse primer:
5’-CCCCACCCTAGGAGAACTTCTCTTT-3’
The following primers are used for the D1S80 VNTRThe forward primer:
5’-GAAACTGGCCTCCAAACACTGCCCGCCG-3’
The reverse primer:
5’-GTCTTGTTGGAGATGCACGTGCCCCTTGC-3’
40
It is important for you to pipet accurately and to carry out these experiments carefully
or your results will not be reproducible and definitive – not very good evidence for a
crime scene investigation.
4.1
Part A: To Make Template DNA:
1. You will be assigned a number by the TA. This is your sample identification for
this lab.
2. Label a 50ml tube containing 0.9% saline with your number.
3. Pour all of the saline solution into your mouth and swish vigorously for 60
seconds or more.
4. Return saline to the tube and replace cap.
5. Load your tube into the IEC Centra and centrifuge at 2,800 rpm for 10 minutes.
6. Carefully decant the supernatant into the sink; do not disturb the pellet. Place
the tube on ice.
7. Resuspend the 10% Chelex solution and add 500 L to the pellet.
Note: When dealing with Chelex make sure to use the wide-mouth pipette tips.
8. Transfer 500 L of the resuspended Chelex/cell solution to a sterile 1.5ml
microfuge tube labeled with your name.
9. Incubate your sample in a boiling water bath for 10 minutes.
10. Remove your sample from the water bath and cool on ice for ~1 minute.
11. Spin your sample in the microcentrifuge for 30 seconds to pellet the Chelex
beads.
12. Transfer 200 L of the supernatant to a clean, sterile 1.5 ml tube labeled with
your number and place it on ice. Do not transfer any of the Chelex.
13. Waste is biohazardous! Discard appropriately.
Part B: Set Up the PCR:
1. Label the cap of a 0.5 ml PCR tube with your initials.
2. Use the amplification chart on the following page as a checklist while adding
reagents to the 0.5 ml PCR tube. Do not cross contaminate! Note: Keep all your
tubes on ice while adding materials to them. Keep them on ice until you are
ready to load into the thermocycler.
3. Add the following reagents accurately:
Reagents (uL)
Tube 1 TPA-25
Template DNA
5
10X PCR buffer
5
25 mM MgCl2
5
Tube 2 TPA-25
5
5
5
Tube 3 D1S80
5
5
5
Tube 4 D1S80
5
5
5
41
1.25 mM dNTPs
8
Forward Primer
1
Reverse Primer
1
Taq polymerase
0.5
Sterile dH2O
24.5
(total volume/tube= 50 ul)
8
XXX
1
0.5
24.5
8
1
1
0.5
25.5
8
1
XXX
0.5
25.5
Make sure you put the correct primers in their corresponding tubes! You will be
given 4 different primers: Forward & Reverse for D1S80 and Forward & Reverse for
TPA-25.
4. Cap your tubes and place them in the thermocycler. Select the appropriate
protocol for each set of primers.
5. After preparing the tubes for the PCR, store your remaining DNA in a labeled
tube at -20C.
TPA-25 PCR Conditions
94C
94C
56C
72C
72C
4C
5 min
1 min
1 min
1 min
10 min
HOLD
Initial Denature
30X
Final Elongation
****************
D1S80 PCR Conditions
94C
94C
63C
72C
5 min
1 min
1 min
1 min
94C
1 min
Initial Denature
5X
42
64C
72C
1 min
1 min
25X
72C
4C
10 min
HOLD
Final Elongation
****************
4.2 Preparation of 2% Agarose Gel
WEAR GLOVES WHEN PREPARING AND RUNNING GELS!
1. Prepare a 2% agarose gel.
2. Microwave for ~1.5 minutes to dissolve the agarose.
3. Let the agarose cool slightly before pouring, ~2 minutes. Note: The 2% agarose
solution will solidify quickly! Pour gel while still relatively hot.
4. Pour the agarose into the gel-casting tray.
5. While the gel is setting, prepare your DNA samples.
6. In a 0.5ml tube, mix 4µL sample buffer with 20µL unamplified DNA. Repeat
with each PCR product.
7. Obtain an aliquot of 100bp ladder from the Lab Tech; this will be loaded directly
on the gel.
8. Once your gel is set, remove the comb and place the gel in the running box.
9. Cover the gel with TAE buffer and load your DNA samples.
10. Run gel at 120 volts for 45 minutes to 1 hour.
11. Once the electrophoresis is complete, stain your gel for ~10 minutes in ethidium
bromide. WEAR GLOVES! Ethidium bromide is a mutagen and carcinogen.
12. Examine, photograph, and destain gel.
4.3 Interpreting your results_____________________________________
Examine your results and consult additional literature. Draw preliminary conclusions.
Your TA will ask you to describe and explain your gel.
4.4 to 4.5 Repeating PCR
When dealing with forensic science, tests need to be repeated to make sure the results
are the same and therefore correct. In 4.1 and 4.2, you isolated your own DNA, used
PCR to amplify it, and ran it on a gel. Now, you will use the remaining isolated DNA
and repeat PCR using varying amounts of water, MgCl2, dNTPs, primers, template
43
DNA, 10X PCR buffer, and Taq. Use your past knowledge and experience of PCR to
calculate the amounts and concentrations of the PCR reagents to use.
4.6 to 4.7 Murder Investigation
You are a lab technician working at a forensics lab. A DNA sample has come in. The
sample is evidence that has come from a murder scene and the murderer is unknown.
The DNA sample was collected from a bottle found at the crime scene. The
investigators have swabbed the bottle and placed the DNA in a sterile saline solution to
preserve it until processing. Investigators have also collected DNA samples from 4
murder suspects and from the murder victim. You must process and compare the DNA
sample found at the crime scene to those collected from the murder suspects and the
victim. From your results, determine who is guilty of the crime and prove why they are
guilty.
In addition, other forensics labs are processing the same evidence/samples. Your results
should match up with all other labs if they are correct. If your results do not match up
with the other forensics labs’ results, trouble shoot and repeat until all labs have the
same results. Follow the protocols that you have just used with your own DNA to
prepare the DNA samples for PCR.
44
Module 5 Microarray Technology
Introduction
Modified from Microarray Technology Lab Manual by Ahmad Chaudhry, et al. Vermont Genetics Network,
Microarray Outreach Program, University of Vermont, Spring 2004.
In this module, students will learn how gene expression in yeast (Schizosaccharomyces
pombe) is changed after exposure to oxidative stress caused by the addition of
hydrogen peroxide (H2O2), an oxidizing agent. Experimentally, the yeast will be
grown for 48 hours in 1/2x YPD broth containing 3x glucose. The culture will be split
into a control and treated group. The control will be exposed to Hanks Buffered Saline
Salt (HBSS) only, while the treated yeast are exposed to 0.5mM H2O2 for 1 hour. RNA
will be extracted and prepared for use on Affymetrix® GeneChips. The final synthesis
product is taken back to the UVM Microarray Core Facility and hybridized to the
yeast GeneChips. The resulting gene expression data will be uploaded into special
bioinformatic data analysis software where students are taken through complex data
analysis procedures.
Days One:
Today we will be treating the yeast Schizosaccharomyces pombe (a haploid fission
yeast) [NRRL Y-128 or ATCC38366] with hydrogen peroxide [H2O2] to understand
the gene expression effects during oxidative stress conditions in this simple
eukaryote. The H2O2 treated yeast, and an untreated control will be lysed using a
combined procedure employing a lyticase enzyme and a buffer containing
guanidinium isothiocyanate after the 1-hour treatment procedure. The first step uses
lyticase which degrades the poly- -1,3 glucan bonds in the cell wall of fungi,
resulting in the formation of spheroplasts (a fungal cell without a cell wall). The
second step uses RLT buffer (guanidinium isothiocyanate) that ruptures the cell
membrane thereby allowing recovery of total RNA on a common silica gel spin
column manufactured by Qiagen.
The Silica Column Chemistry
Guanidium isothiocyanate (GITC or GSCN) is a chaotropic salt at about 5 M in the
RLT buffer. This solution at a reduced pH (6.4) and used in conjunction with ethanol
and water disrupts the molecular characteristics of water and forces the RNA to be less
soluble. This promotes nucleic acids to electrostatically bind to the silica. A
stoichiometric balance of GITC, ethanol, pH, and water controls the binding of either
RNA or DNA to the silica. Therefore, changes in this stoichiometry will select a
majority of either one or the other. This not a 100% selective procedure and DNase
45
treatment is required. The use of ethanol in rinse RPE buffer maintains the bond
between the silica and RNA. The recovery of RNA off the membrane is accomplished
with water because RNA is very soluble in water and can no longer maintain an
electrostatic bond with the silica once the water structure has been re-established.
Day Two:
We will be preparing the mRNA for microarray target preparation and then, cDNA
synthesis.
Technical Overview:
Today you will be starting to prepare the target for the Genechip analysis. This is a
lengthy procedure and will be broken down over several lab periods. In short, you
will be starting with mRNA, which comprises 0.5-2% of the total RNA in typical
eukaryotic cells. First, you will generate the first complementary DNA (cDNA)
strand from the mRNA. This is called the first strand cDNA synthesis. This step
utilizes a primer with a sequence of 24 T's in a row and is called oligo d(T), which
binds to the poly A tail of eukaryotic mRNA. The primer also contains the consensusbinding site for T7 RNA polymerase (T7 promoter sequence). Once the primers have
been hybridized to the poly-A mRNA, reverse transcriptase is added and the first
strand of cDNA is synthesized. It is important to note that the 5s, 5.8s, 18s, and 25s
rRNA do not have a poly-A tail and will not be primed or synthesized in this procedure.
46
The sequence of T7 Oligo d(T)24 is:
T7
d(T)24-Binds
to PolyA of mRNA
The next step in the cDNA reaction is the synthesis of the second cDNA strand from
the first cDNA strand. This is done by adding RNase H, DNA polymerase I, and
DNA ligase. The RNase H causes many strand nicks in the phosphodiester bonds of
the bound mRNA strand creating the availability of 3'OH groups. DNA polymerase
uses these 3’ OH groups as a priming site; thus synthesizing another cDNA strand
until it reaches another nicked site. The DNA polymerase can only displace RNA,
not DNA. After cDNA has been synthesized from all the nicks, the areas between the
nicks need to be filled in or ligated. This is accomplished using DNA ligase. After
the two hour synthesis process, all the RNA has been replaced with cDNA and
ligated to form one large cDNA strand. However, because the DNA polymerase
does not synthesize completely to the end of the strand, a T4 DNA polymerase is
added. This ensures the synthesis of the T7 promoter site from the original T7 oligo
d(T) primer used in the first synthesis reaction.
Synthesis of cDNA:
The starting material for your target preparation will be high quality, fully intact,
total RNA from yeast. The minimum concentration to start with is 100-270 ng/µl.
After isolation and purification of the yeast RNA, the concentration is determined
by absorbance at 260 nm on a spectrophotometer (1 O.D.= 40 µg/mL RNA). The
A260/A280 ratio should be approximately 1.8-2.1. We are checking the quality of
the RNA by running an aliquot on the Agilent Bioanalyzer 2100 (performed prior to
class at UVM core) and running on an agarose gel (EGEL) prior to starting the
assay. More appropriately, a denaturing formaldehyde gel would be used, but we
will not have time to perform this.
47
Overview of cDNA Synthesis Reactions
48
Day Three:
We will be cleaning the cDNA and preparing it for an in Vitro Transcription to
synthesize an antisense cRNA.
Cleaning the cDNA:
After synthesis of the cDNA, it will need to be purified before proceeding to
the next step. This will be accomplished using a solution of phenolchloroform-isoamyl alcohol (PCI) at a specific pH. The underlying principle is
that both phenol and chloroform cause proteins/enzymes to become denatured
and subsequently soluble in the organic phase, while the cDNA will remain in
the aqueous phase. This is performed by mixing the cDNA sample with PCI
and transferring the full volume to a phase lock gel tube. The phase lock gel
tube is centrifuged allowing separation of the aqueous and organic phases
with a gel barrier. This allows for easy recovery of the cDNA in the aqueous
phase. It is important to note that the pH of the PCI is important because this
will determine which layer the cDNA will be retained in (the aqueous or
organic phase). To ensure the cDNA is in the aqueous phase, the pH of the
PCI needs to be basic, which is accomplished through a layer of TRIS pH 8.3
over the PCI solution.
The aqueous phase is further purified through a precipitation step using
ethanol [an alcohol] and ammonium acetate [a salt]. Because DNA is negatively
charged, salt is used to mask the charge, causing it to precipitate. The ethanol is
used in conjunction with the salt because it is non-polar and DNA will not
readily dissolve in it. This reaction is sometimes done at -20°C to encourage
maximum DNA precipitation. After the precipitation, the cDNA is centrifuged
to a pellet with a visualization agent called Pellet Paint, an additive that
contains glycogen (a co-precipitate) and a dye. This helps form a high quality
visible pellet on the bottom of the tube. The pellet is then washed several
times with 80% ethanol, dried, and resuspended in DEPC water.
The In vitro Transcription (IVT):
The IVT step uses the T7 promoter site that is attached to the T7 Oligo d(T)24
primer used in the first strand cDNA synthesis. A T7 RNA polymerase binds
to this site and copies the opposing cDNA strand, synthesizing an antisense
cRNA using the two standard nucleotides, A and G and two modified
nucleotides; biotinylated uracil and cytosine. This generates a complementary
cRNA [antisense] strand to the original mRNA with biotinylated nucleotides.
49
These biotinylated nucleotides will be needed in the next step for staining of
the RNA using strepavidin phycoerytherin.
Day Four:
We will be cleaning the cRNA before fragmentation. It is important to have
“clean” cRNA before applying to the chip to reduce non- specific binding, as well
as being sure that the cRNA has fragmented.
Cleaning and fragmenting the biotinylated cRNA:
Using the Qiagen RNeasy system, the biotinylated cRNA must be purified. The
cRNA needs to be free of enzymes, dNTP's, reducing agents such as DTT
[dithiothreitol] and resuspended in DEPC water. The clean cRNA can then only
be accurately quantified. Remember that enzymes and dNTP's also have an
absorbance at 260 and/or 280nm that will affect the quantitation of the molecµle
of interest. That is why [for instance] you cannot quantify a raw PCR reaction
before and after thermocycling, because all components are still there, just
assembled differently.
The cleaning is performed by adding beta-mercaptoethanol, ethanol, and a
buffer to the cRNA mix to reduce the proteins. These reagents aid in the
solubilization of proteins and prepares the cRNA for binding to a silica-based
spin column (pink basket of the RNeasy kit). The cRNA is washed and eluted
using DEPC water. The cRNA is then quantified using a spectrophotometer at
260nm for nucleic acids and 280nm for proteins. A 260/280nm ratio of 1.8-2.1
indicates a clean cRNA sample that can be used for subsequent steps. 260/280
ratios of less than 1.8 indicate that the cRNA is contaminated with too much
protein and should be purified again.
10ug
30’
50
Performed by UVM:
Once the cRNA is cleaned and quantified, it will be given to UVM’s
Micoarray Core Lab to perform the following last steps. The hybridization
mixture contains several controls that provide information about the success of
the hybridization procedure. These controls are the B2 oligo for which there are
several hundred probes along the outer edge of all expression arrays and
checkerboard pattern in each corner. These predefined patterns provide signals
for the Affymetrix Microarray Suite software to perform automatic grid
alignment during image analysis. They can also be used to align the grid
manually. The fluorescence intensities for control oligo B2 are not used for
analyzing data. The bioB, bioC and bioD are biotinylated gene fragements much
like your sample, which represent the biotin synthesis pathway from the bacteria
E. coli. The Cre control is the recombinase gene from P1 bacteriaphage. These
biological controls are prepared at standardized concentrations and signal
intensities obtained on these genes provide information on how well the
hybridization, washing and staining procedures have performed.
After the hybridization mixture has been made, it is denatured at 94C for
5 minutes and injected into the GeneChip through the injection ports on the back
of the chip (See diagram in appendix). The GeneChip is then placed in a rotating
incubator and the biotinylated target that you prepared is allowed to bind
(hybridize) to the sequence specific probes on the array.
The staining procedure involves using a fluidic station designed to
accommodate GeneChips. Two solutions are prepared: a strepavidinphyoerytherin dye (SAPE) and a biotinylated anti-phycoerytherin antibody
(BAP). Before placing the GeneChip into the fluidic station, the fragmented
cRNA hybridization mixture is manually removed and replaced with buffer.
The chip is placed in the fluidics station and an automatic process of
washing and staining occurs for approximately 2 hours. The procedure starts by
first staining with SAPE. The SAPE will bind the biotinylated cRNA that is
hybridized to the probes on the chip. It is washed and then stained with BAP.
BAP will bind to the SAPE. The chip is washed again and another SAPE staining
is carried out to bind to the BAP. The double staining procedure is employed to
amplify the signal intensities. This allows for the detection of low transcript
mRNA’s that might otherwise not be detected. After staining, the sample is
washed and ready for scanning.
The GeneChip is scanned after the washing and staining protocols are
complete. The chip is placed in the scanner that is equipped with an Argon
488nm laser and the beam is scanned back and forth across the glass surface at a
51
resolution of 5m. Fluorescence is detected using a high-resolution photo
multiplier tube (PMT). This enables high-resolution detection of fluorescence
from the probe set feature (pixel), which is only 19m square. There are ~1
million probe areas on each GeneChip!
After collecting and analyzing the image data, the results are archived and
burned onto a CD. Because there is so much data, it is necessary to analyze it
using a special software package such as GeneSifter. This software can handle
tens of thousands of data points at one time and data analysis can often take
months. Therefore, it is critical that the data meet specific control specifications
before this time consuming analysis is performed.
52
Image from http://www.microarray.lu/en/MICROARRAY_Overview.shtml Microarray Center
CRP
53
TO CREATE AN RNase-FREE ZONE:
**It is very important that you follow these directions exactly—minimizing
contamination and degradation of your samples is crucial to the success of the
experiment**
1. Tape off an area with labeling tape. Label the area as an “RNase-free
zone”.
2. Wearing gloves, wipe down the area with ethanol, then with the RNase
Away® solution.
3. Wipe down the pipette men you will use during your experiment with the
RNase Away® solution.
4. Once the pipette men are cleaned, place them in the RNase-free zone.
5. Wipe down any instruments or bottles that you will come in contact with
(racks, tube holders, etc.) with ethanol. Place them in the RNase-free zone.
Make sure to read through the procedure and plan accordingly.
6. Follow the guidelines bellow to insure an RNase-free zone:
7. If you are to touch any part of your body or hair with your gloved hands,
change your gloves. Your skin and hair naturally contain DNase and
RNase to battle against everyday bacteria and germs.
8. If you walk to another area in the lab, change your gloves before working
in the RNase-free zone again.
9. If you touch a piece of equipment (microcentrifuge) or lab bench that has
not been wiped down, change your gloves before working in the RNasefree zone again.
10. Use only unopened sterilized tips, tubes, etc. If you are unsure about the
“freshness”, ask your Lab Tech.
11. Use only sterile distilled water.
12. Work quickly, but carefully. RNA degrades quickly.
Day Before 5.1: Preparation of Yeast Cultures (Done for you)
Using sterile technique, 500mL of YPD (Yeast Peptone Dextrose) broth will be
inoculated with an isolated yeast colony.
The culture will stir on a stir plate at room temperature (22-25C) for 48 hours.
5.1 Treatment of Control and Treated Yeast Cultures (DAY 1)
1.5 Hours Before Lab (Done for you):
1. From the 48 hour old culture, aseptically transfer 40.0 ml of the broth
culture into two (2) sterile 125 ml flasks each containing a stir bar using a
sterile 50 ml pipet.
2. Label one flask as “treated” and aseptically add enough H2O2 from the
“working stock" to achieve a final H2O2 of 0.5mM [See below]. Label the
54
control flask as “control” and add an equal amount of HBSS as you did
H2O2.
Preparing and Determining the Concentration of H2O2
a. Preparation of H2O2 Stock Solution: Combine 5.0µL of 30% H2O2
and 495 µL of Hanks Balanced Saline Salt [HB without phenol red,
Mg, or Ca. Vortex.
b. Working H2O2 solution : Make a 1:10 dilution by combining
100µL H2O2 stock solution with 900µL HBSS and vortex. Use this
for spectrophotometer measurement and for the experiment.
c. Blank the spectrophotometer with HBSS
d. Measure the absorbance at 240nm.
e. Concentration of H2O2 in Working Solution : (Ab240) x 229 =
____mM of H2O2 [B].
f. Example: H2O2 to add to 40 ml flask= A/B x 40ml x1000 = ul of
working solution to use in experiment
Where A=0.5mM B=your concentration in working solution
3. Place each flask on separate stir plates and stir at room temperature for 1
hour at [as close as] the same speed as possible. Cultures should be ready
for harvest 30 min after the start of the first class period. Therefore, H2O2 is
added 30 minutes before the start of the class.
During Lab:
1. Prepare an RNase-free work zone according to the directions above..
2. Prepare fresh lytic enzyme at 10 Units/µl by adding 1 ml of DEPC water
directly to the lyticase vial.
a. 10,000 U Lyticase
b. 1 ml DEPC Water
3. Vortex well and invert several times to insure complete mixing.
4. Transfer to a 1.5mL microcentrifuge tube. This solution is stable for 12
hours.
5. Prepare fresh DNase I solution using the Qiagen DNase kit and store on
ice.
6. Add 10L DNase I and 70L RDD Buffer to a 1.5mL microcentrifuge tube
7. Transfer 1.5mL of the Control yeast culture to a 1.5mL microcentrifuge
tube.
8. Transfer 1.5mL of the Treated yeast culture to a 1.5mL microcentrifuge
tube.
***Perform the following steps 9 through 39 to each yeast culture!
55
9. Spin down 1.5 ml of yeast culture at 8000x g for 2 minutes using an Axygen
1.7 ml tube in a microcentrifuge at room temperature.
10. Carefully remove supernatant (without disturbing pellet) using a P1000
micropipet, discard supernatant
11. Add 1 ml of sterile DEPC water, vortex, and spin again at 8000 x g for 2
minutes.
12. Remove and discard supernatant. Remove as much of the liquid as
possible from the yeast pellet. You may need to use a smaller pipet to get
any remaining liquid.
13. Add the following to the yeast pellet and
gently vortex:
a. SG buffer
100 µl
b. Lyticase solution (10U/µl)
30 µl
14. Incubate for 30 min at room temp (22-25°C), gently swirl tube every 10
minutes to generate spheroplasts. Spheroplasts must be handled
gently.
15. Prepare RLT buffer (from Qiagen RNeasy Kit) for the nest step by adding
10ul of β- mercaptonethanol (BME) to 1ml RLT buffer. USE BME under
fume hood! This buffer will now be called b-RLT.
16. After 30 minute incubation, add 350 µl b-RLT buffer to lyse spheroplasts.
17. Vortex vigorously for 1 minute. Ensure your tube is tightly capped by
holding lid closed while vortexing.
18. Add 250 µl of 100% ethanol to the tube and briefly vortex. Do not
centrifuge. A precipitate may form after the addition of ethanol, but
this will not affect the RNeasy procedure.
19. Apply the entire sample to an RNeasy mini column.
20. Close the tube gently and centrifuge for 15 sec at full speed.
21. Discard the flow through tube and place the RNeasy column into a new
2 ml capture tube.
22. Add 350 µl RW1 buffer to the RNeasy column to wash the column.
Close the tube gently and Centrifuge for 15 sec at full speed.
23. Apply 80 µl of DNase I solution to the middle of the Qiagen column
membrane. Incubate at room temp for 15 min.
24. Transfer RNeasy column to a new 2 ml capture tube.
25. After the 15 minute incubation, add 350 µl RW1 buffer to the RNeasy
column and spin at full speed for 15 seconds.
26. Discard the flow through tube and place the RNeasy column into a new 2
ml capture tube.
27. Pipet 500 µl RPE buffer onto the RNeasy column to wash the column.
Close the tube gently and centrifuge for 15 seconds at full speed.
28. Discard the flow through tube and place the RNeasy column into a new 2
ml capture tube.
56
29. Add another 500 µl RPE buffer to the RNeasy column. Close the tube
gently and centrifuge for 15 sec at full speed.
30. Place the RNeasy column in a new 2 ml capture tube and
centrifuge in a microcentrifuge at full speed for 1 minute to “dry” the
silica membrane.
31. To recover the RNA, transfer the RNeasy column to a new 1.7 ml
microcentrifuge tube.
32. Pipet 30 µl of DEPC water directly onto the very center of the
RNeasy silica-gel membrane. DO NOT TOUCH THE SILICA GEL
MEMBRANE. Look closely as you perform this step. Use both hands
when pipeting, one on top and one down by the tip to guide the
pipet. Make sure the water was evenly distributed on the
membrane.
33. After allowing the water to incubate on the membrane at room
temp for 1 minute
34. Centrifuge at full speed for 30 seconds.
35. Carefully remove the 30 µl which is recovered in the 1.7 ml tube and
pipet it back onto the center of the silica membrane of the same column.
36. Place column back in to the same 1.7 ml tube and spin again for 1 minute
at full speed. This double elution ensures that the entire membrane was
extracted.
37. Transfer the recovered RNA to a new 1.7 ml tube and label it with
date, sample name, and what it is. Write clearly!!
38. Transfer 4 µl of RNA to a new tube for transport back to UVM for
Nanodrop quantification and RNA assessment. Label this tube carefully
with date, name, and RNA.
39. Keep the samples on ice and quantify samples using the
spectrophotometer at a 1 to 50 dilution (1µl sample + 49µl H2O). This
will be demonstrated by the instructor.
[Sample concentration:_________ ]
[Write this on your tube]
40. Evaluate the RNA by agarose gel electrophoresis . This will be done as a
class. Pre-run the gel for 2 minutes and turn off. Add 14 µl of water to
each well on the E-gel. Add 1µl of each sample to each well and pipet
up and down to mix. Run the gel for 20 minutes.
**Note: If no E-gel then proceede with section 5.2**
41. Samples should be labeled properly for return to UVM and analyzed
using the Agilent Bioanalyzer 2100. This instrument is used to determine
57
the condition of the RNA. Only fully intact RNA is eligible for microarray
analysis. Why?
5.2 Quantifying & Evaluating Samples
1. Thaw your RNA samples on ice.
2. Quantify samples using the spectrophotometer at a 1 to 50 dilution
(1L sample + 49L sterile dH2O). Read at
nm.
Sample concentrations:
Control: ___________
Treated: ___________
3. Prepare a 1.2% agarose gel.
4. Mix 200L of sterile dH2O with 3L of 6X loading dye and mix.
5. Add 1L of your sample to the water/dye mix.
6. Load 20L to each well.
7. Load 20L of the 0.24-9.5Kb RNA ladder.
8. Run the gel for 60 minutes at 100 Volts.
9. Store your remaining samples at -20C.
10. Stain your gel with Ethidium Bromide, visualize with UV light,
photograph and record results.
Note: Ethidium Bromide is a carcinogen and mutagen. Wear gloves!
11. Your samples will be analyzed using the Agilent Bioanalyzer 2100 to
determine the condition of the RNA. Only fully intact RNA is eligible for
Microarray analysis. See Appedix for sample bioanalyzer data.
5.3 Microarray Target Preparation; 1st Strand cDNA Synthesis (DAY
2)
Synthesis of double-stranded cDNA from total RNA is performed by priming off
the poly-A tail of mRNA using T7 Oligo (d)T24. It is important to note that the 5s,
5.8s, 18s and 28s rRNA do not have a poly-A tail and will not be synthesized in
this procedure. The sequence of T7 Oligo (d)T24 is:
5’ – [GGCCAGTGAATTGTAATACGACTCACTATAGGGAGGCGG-(d)T24] – 3’
Note: Spin all reagent tubes before starting as some have only 1 μl in
them!!!!
1. Adjust 1-3 µg (determined by instructor) of RNA to 11µl with DEPC water in a
RNase-free 0.5ml tube. Keep the tube on ice.
58
2. Primer Hybridization: Combine the following reagents:
a. T7 oligo d(T)24
2μl
b. RNA(1-3 μg) 10μl
3. Vortex, and spin in microcentrifuge [full speed] for 5 seconds.
4. Put in a thermocycler at 70°C for 10 minutes.
5. While the 70°C incubation is in progress prepare the following
master mix. Add the following reagents IN ORDER to make the first
strand master mix.
a. First Strand Buffer 5X
4 μl
b. 0.1M DTT
2 μl
c. 10mM dNTP
1 μl
d. Superscript II
1 μl
6. Vortex and spin in microcentrifuge [full speed] for 5 second and place on
ice.
7. After the 70°C step, add the master mix to the RNA primer tube and
incubate in a thermocycler at 42°C for 60 minutes.
8. After the incubation, place on ice. During this incubation, prepare the second
strand master mix.
Second Strand cDNA Synthesis
9. Make the following master mix in a separate tube. Keep it on ice. All tubes
may need to be centrifuged to get the contents to the bottom of the tube
because some tubes only contain 2ul of enzyme!
a. DEPC Water 91 µl
b. 5x Second Strand Buffer
30 µl dNTP(10mM)
c. 3 µl E.coli DNA Ligase (10U/µl)
1 µl
d. E.coli DNA Polymerase I (10U/µl)
4 µl
e. E.coli RNase H (2U/µl)
1 µl
Total volume to be added to First Strand tube
130 µl
10. Vortex master mix and perform a 5 second quick spin in the
microcentrifuge [full speed.
11. Add this 130 µl second strand master mix to first strand tube after the 42C
incubation.
12. Vortex and perform a 5 second quick spin.
13. Incubate for 2 hours at 16°C in thermal cycler.
14. At the end of 2 hours and while the sample is still at 16°C, add 2µl of T4
DNA polymerase and incubate for exactly 5 minutes. Mix well. Do not
incubate longer then 5 minutes or the quality of the cDNA may decrease
59
due to the 3’ to 5’ exonuclease activity of the polymerase.
15. At the end of the 5 minute incubation, add 10 µl of 0.5 M EDTA to stop
the reaction.
16. Store sample at -20°C.
Primer Hybridization:
Combine 1L of T7 oligo dT and 11L of RNA (3g) in a 1.5mL microcentrifuge
tube.
Vortex and briefly spin at 4000rpm. Place in a thermocycler set at 70C for 10
minutes. Place on ice immediately after.
17. Combine 1L of T7 oligo dT and 11L of RNA (3g) in a 1.5mL
microcentrifuge tube.
18. Vortex and briefly spin at 4000rpm.
19. Place in a thermocycler set at 70C for 10 minutes. Place on ice
immediately after.
20. Add the following reagents to a 1.5mL microcentrifuge tube:
a. 1st Strand Buffer 5X
4L
b. 0.1M DTT
2L
c. 10mM dNTP
1L
Vortex and briefly spin at 4000rpm. Place in a thermocycler set at 42C for 2
minutes.
21. Add 1L of Reverse Transcriptase (Superscript II).
22. Vortex and briefly spin at 4000rpm.
23. Place in a thermocycler set at 42C for 60 minutes.
24. After the incubation, appropriately label your tubes and store at -20C.
5.4 Precipitating the cDNA, Cleaning the cDNA, and Setting Up the
In Vitro Transcription (IVT)
(DAY 3)
Note:
Phenol Chloroform Isoamyl Alcohol is at a ratio of 25:24:1 and saturated with
10mM Tris-HCl pH 8. Phenol is an organic acid and care must be used when
handling. Phenol can burn skin, be absorbed into the circulatory system and
cause health problems. Chloroform is an organic solvent and should be used in
the hood.
Phenol
– Corrosive
– Local anesthetic
60
– Can severely damage eyes and skin
– Can cause severe burns of the skin and eye
– Poisonous if ingested-can be fatal
Chloroform
– Can cause irreversible corneal injury
– Fatal if swallowed
– Flammable
Isoamyl Alcohol
– Flammable
– Poisonous
Precipitating the cDNA:
1. Centrifuge a Phase Lock Gel tube at full speed for one minute. DO NOT
VORTEX.
2. Add 162 µl of the bottom layer from the Phenol Chloroform Isoamyl
Alcohol (PCI) to the cDNA synthesis reaction and vortex for 2 seconds.
Hold cap tightly as leaking can occur during the vortex.
Note: PCI tends to leak out of the pipet so work quickly.
3. Transfer the cDNA-PCI mixture to the phase lock gel tube. DO NOT
VORTEX the phase lock gel tube.
4. Centrifuge at full speed for 2 minutes.
5. Transfer the top layer to a 1.7 ml microcentrifuge tube.
6. Add the following to the 1.7 ml microcentrifuge tube and vortex.
Ethanol (100%)
405 µl
NH4OAc (7.5M)
80 µl
Pellet Paint
1 µl
7. Centrifuge at 12000 x g [full speed] for 20 minutes at room temp.
8. GENTLY remove the tube from the Centrifuge being careful not to
disturb the cDNA pellet. The pellet should be pink and approximately the
size of a grain of salt. Put on ice and immediately proceed to next step.
Keep your tube on ice.
**At this point a tiny pink visible pellet should be present on the
bottom of the tube. If not, see the instructor.**
61
Cleaning the cDNA Pellet:
9. Using a micropipet, carefully remove the liquid from the tube being very
careful not to disturb the pellet. Tip the tube to enable removal of as much
liquid as possible. Remember that the pellet is your sample!
10. Add 500µl of ice cold [-20C] 80% ethanol to the tube with the pellet.
Gently cap tube and invert slowly several times. Watch your pellet very
closely. Place tube back in rack and let the pellet settle to the bottom of
the tube for a minute. Alternately, you may Centrifuge the tube at full
speed for 15 seconds to get the pellet back down to the bottom of the
tube. [See video on website for this technique]
11. Using a 1 ml micropipet, carefully remove the ethanol being very careful
not to disturb the pellet. Tip the tube to enable removal of as much liquid
as possible.
12. Repeat steps 2 and 3 with a new aliquot of 80% ethanol.
13. Finally, remove all of the ethanol. Centrifuge the tube again [full speed]
for about 5 seconds and using a smaller pipet such as a P20 or P200,
remove the last few microliters. The goal is to remove as much ethanol
as possible without disturbing you pelleted cDNA.
14. Place the tube with the pellet in a drying box for 10 minutes to evaporate
all of the ethanol. The dried pellet is easily lost once it is dry. Be very
careful to handle the tube gently. Close the cap gently. Visualize the
dried pellet to confirm it is present in the tube.
15. Resuspend the pellet in 22 µl of DEPC water and put on ice.
In Vitro Transcription (IVT): Synthesis of biotin labeled cRNA from cDNA
using the Enzo Kit:
16. Using the ENZO bioarray kit, a master mix for the entire class will be
prepared as follows. The instructor will prepare this mix or designate
someone from the class. This must be done an RNase-free area free from air
currents.
62
Amt/sample #Samples Total
a. Reagent 1 [10x Reaction buffer]
4 µl
b. Reagent 2 [10x Biotin nucleotides] 4 µl
c. Reagent 3 [10x DTT]
4 µl
d. Reagent 4 [10x RNase Inhibitor]
4 µl
e. Reagent 5 [20x T7 RNA polymerase]
2 µl
Total Volume
18 µl
NOTE: Be sure to make extra for this step. Add enoµgh for 1 more reaction
17. Combine the following in a 0.5 ml microcentrifuge tube and pipet up and
down several times to mix. Spin in Centrifuge full speed for 5 seconds.
a. Clean cDNA
22 µl
b. Enzo master mix [from above]
18 µl
Total Volume
40 µl
18. Incubate the above mixture at 37°C for 16 hours in the thermocycler.
19. Store the sample at -20 C after the 16 hour incubation is complete (to be
done by instructor).
5.5 Cleaning the Biotinylated cRNA Using the Qiagen RNeasy Kit
and Fragmentation
(DAY 4)
Note: Add 100L BME per 1mL RLT Buffer. RLT Buffer is stable for 1 month
after addition of BME. We shall call this reagent b-RLT. All steps of the RNeasy
protocol should be performed at room temperature.
1. Transfer the cRNA sample to a 1.7 ml microcentrifuge tube and add 60 µl of
DEPC water and 350 µl BME-RLT and vortex.
2. Add 250 µl ethanol (96–100%) and lightly vortex again.
3. Apply the sample (700 µl) to an RNeasy mini column placed in a 2 ml
collection tube (supplied).
4. Close the tube gently and Centrifuge for 15 s at full speed.
5. Remove the pass-through liquid from the tube and reapply it to the same
RNeasy mini column again.
6. Close the tube gently and Centrifuge for 15 s full speed. Transfer the
RNeasy column into a new 2 ml collection tube and discard the flowthrough and collection tube.
7. Pipet 500 µl of RPE buffer onto the RNeasy column. Close the tube
gently, and Centrifuge for 15 s at full speed to wash the column.
8. Transfer the RNeasy column into a new 2 ml collection tube. Discard the
flow-through and collection tube.
63
9. Add another 500 µl RPE buffer to the RNeasy column. Close the tube
gently and Centrifuge for 15 s at full speed to dry the RNeasy silica-gel
membrane.
10.Transfer the RNeasy column into a new 2 ml collection tube and
Centrifuge in a microcentrifuge at full speed for 1 min.
11.Transfer the RNeasy column into a new RNase-free 1.7 ml
microcentrifuge tube.
12.To recover the RNA from the membrane, pipet 30 µl of DEPC water
onto the RNeasy silica-gel membrane and wait 1 minute. Make sure
the DEPC water contacts the entire membrane. Do not touch the
membrane. Close the tube and Centrifuge for 1 min at full speed to
elute.
13. Remove the eluted RNA and apply it to the membrane again. Close
the tube gently and Centrifuge for 1 min at full speed to elute. This
double elution results in a slightly higher cRNA yield.
14. Transfer the cRNA to a new microcentrifuge tube.
15. Quantify samples using the spectrophotometer.
a. Add 49 µl of water to a Uvette and add exactly 2 µl of sample.
b. Pipet up and down to ensure all cRNA has been expelled (2µl
sample + 48 µl H2O).
c. Use a p200 pipet set to 25ul and mix by pipetting
up and down to mix well. Tap out all bubbles from
Uvette.
d. [Sample concentration:_________ ]
Calculating the Adjusted cRNA Concentration
Calculation of the adjusted cRNA concentration must be done in order to
subtract out the original untranscribed RNA [i.e. tRNA, 5s, 18s, and 25s rRNA].
The cRNA is fragmented using a metal-induced hydrolysis reaction to create
cRNA fragments of 35-250 bp to be used in hybridization to the yeast GeneChip.
Both the unfragmented and fragmented cRNA are visualized on an agarose gel.
This is important because we need to ensure that the unfragmented cRNA is
not degraded from contamination during the handling steps leading up to this
point.
16.Calculate the Adjusted cRNA concentration using the following formµla:
a. ADJ cRNA= {Amount of clean cRNA} - {Total amt RNA used at
start}
64
b. Example: 52.9μg –2μg=50.9μg or 50,900ng
c. A) What is the concentration of your cRNA in ug [not ng] per μl:
__________ ug/μl
d. B) How many μl do you have:__________μl
e. C) How many μg of total RNA did use on day 1:__________μg
f. [AxB]-C = adjusted cRNA concentration
Fragmenting the cRNA for Target Preparation
Affymetrix recommends that the cRNA used in the fragmentation procedure
be sufficiently concentrated to maintain a small volume during the
procedure. This will minimize the amount of magnesium in the final
hybridization cocktail. The cRNA must be at a minimum concentration of 600
ng/µL. The fragmentation buffer contains a basic solution of TRIS acetate,
magnesium acetate, and potassium acetate which causes a metal induced RNA
hydrolysis at high temperature.
17.Setup the following fragmentation reaction in a 0.5 ml Centrifuge tube. Use
a 0.5 ml tube for this reaction
Adj cRNA concentration: __________ ng/μl
10μg=___________μl
Volume
Volume (μl)
10 μg of adj cRNA
1-16 μl
5x fragmentation buffer
4 μl
DEPC water
(as needed to make the final volume 20 μl)
Total reaction volume
20 µl
20 µl
18.Vortex and Centrifuge briefly.
19.Incubate at 94°C for 30 minutes in a thermocycler. Put on ice
following the incubation.
Fragmentation: Hammerhead Cleavage
Lets look at the molecμlar level
Fragmentation buffer
94C for 30 minutes
20.
50-200 bp fragments
65
The cyclic nucleotide is now a 2’-nucleotide.
Hydrolysis at the 2’ carbon produces the 3’nucleotide.
Evaluating The Samples on a Standard Agarose Gel
This procedure will evaluate your unfragmented and fragmented cRNA. Samples
will be loaded on a precast E-GEL and allowed to run for 20 minutes.
23.Remove a E-Gel from pouch and place on E-Gel base.
The intermediate product is
24.Pre-run the gel for 2 minutes by pressing the button continuously
for 10
a 2’, 3’ cyclic nucleotide.
seconds. You will see the light start to blink indicating it
is next
in a step
pre-run
The
is hydrolysis
at
the
3’
carbon.
mode. It will beep and stop after 2 minutes.
25.Add 15 µl of water to each well on the E-gel.
26.Add 2µl of each sample to each well and pipet up and down to mix well.
Run both the fragmented and unfragmented cRNA.
27. Load 4 µl of Ladder
28. Run the gel for 20 minutes.
29. Visualize the E-gel on a transilluminator.
30. Take pictures.
Note: The fragmented sample should be a smeared band at the bottom
of the gel indicating fragments of RNA between 35 and 250bp. The
unfragmented sample should be a smear from the top to the bottom of the gel.
66
The total RNA [if you run them] should be two bands at the middle of the gel
indicating the 25s and 18s ribosomal RNA units. See picture below.
ladder
2000, 1200, 800, 400, 200, 100bp
The remainder of the experiment will be performed by the UVM
DNA/Microarray Processing Facility. See p. 35 for a detailed summary of what
procedures will be used to complete this experiment.
Day 5: Tour of Core Facility
Day 6: Tutorial of Bioinformatics with Dr. Janet Murray
67
Appendix A: Solutions Guide
SOLUTIONS GUIDE
0.5M NaOH/0.8M NaCl
p. 73
0.5M Tris/ 1.5M NaCl
p. 73
2.0M Tris Cl (pH 8.0)
p. 69
Amido Black Destain
p. 78
Amido Black Stain
p. 78
Ampicillin Stock
p. 69
Blocking Buffer
p. 77
Block. Buf. (0.1%(w/v) BSA) p. 79
Buffer 1
p. 75
Buffer 2
p. 75
Buffer 3
p. 75
CaCl2 (50 mM)
p. 72
Chelex Extraction Solution p. 80
Chloroform:Isoamyl Alc. (24:1)p. 72
Coomassie Blue Destain
p. 77
Coomassie Blue Stain
p. 77
Cracking Buffer
p. 73
Denhardt’s (100X)
p. 74
dNTPs mix
p. 76
Heat-treated RNase A
p. 71
Hybridization Solution
p. 74
LB amp
p. 69
Lysis Buffer
p. 76
PAGE Running Buffer
p. 76
Phenol/Chloroform/Isoamyl p. 80
PBS (Phosphate Buffer)
p. 71
Potassium Acetate (5M)
p. 71
Prehybridization Solution p. 74
Primary Antibody Dilution p. 78
Primer Dilutions
Resolving Buffer (4X)
SA-AP
Sample Buffer (6X- for DNA)
SDS Sample Buf. (6X -for pro.)
Secondary Antibody
SG Buffer
Sodium Acetate (3M)
Solution I
Solution II
SSC (20X)
SSC 0.16X /0.1%(w/v) SDS
SSC 0.2X /0.1% (w/v) SDS
SSC 2X /0.1% (w/v) SDS
Stacking Buffer (4X)
Sterile Saline (0.145M)
Sterile Saline (0.9%)
Stop Solution
TAE stock (50X)
TBS (4X)
TBS-T
TE Buffer
Transformation Buffer
Western Blot Transfer Buffer
YPD Nutrient Media
λHind III maker
p. 75
p. 76
p. 75
p. 70
p. 77
p. 78
p. 80
p. 73
p. 71
p. 71
p. 70
p. 75
p. 74
p. 74
p. 76
p. 72
p. 79
p. 79
p. 69/70
p. 70
p. 70
p. 69
p. 72
p. 77
p. 80
p. 70
68
GENERAL SOLUTIONS
Ampicillin Stock
100 mg/ml stock solution: for example, 0.5g ampicillin sodium salt into 5
ml dH20. Filter sterilize, and store at -20°C.
LB amp
100 µg/ml final concentration: 1:1,000 dilution of ampicillin stock into LB
broth. For example, add 1 ml ampicillian stock into 1 L LB broth.
*NOTE: Ampicillin is heat-sensitive, so LB broth must be cooled to 60°C after
coming out of the autoclave, before the ampicillin is added. Setting the water
bath to 60°C and letting the LB broth cool in there for an hour is a good way to
ensure the LB doesn’t solidify (if making plates).
***1ul into 1ml for LB amp***
TE buffer
Need (final conc.):10 mM Tris-Cl (pH 7.5)
1mM EDTA (pH 8.0)
Make: 1.576 g Tris-Cl
0.2922 g EDTA
Bring up to 1 L dH20
2M Tris-Cl (pH 8.0)
177.6g Tris-Cl
10.6g Tris-base
In ~950mL sterile dH20
**Ph 8.0**
Bring up to 1.0L with sterile dH20
50X TAE stock (pH 8.5)
Prepare a Stock Solution of 0.5M EDTA:
93.05g EDTA disodium salt (FW= 372.2)
In ~400ml sterile dH20
***bring pH to 8.0***
Up to 500mL with sterile dH20
69
Prepare a 50X Stock of TAE
242g Tris Base (FW= 121.14)
In ~750ml sterile dH20
Carefully add 57.1mL Glacial Acetic Acid
100mL 0.5M EDTA (pH 8.0)
Bring up to 1L with sterile dH20
pH 8.5, but no adjustment needed
*Dilute 50X stock 1:10 for a 5X stock*
6X DNA sample buffer
Need: 0.25% (w/v) Bromophenol Blue
40% (w/v) Sucrose
Make: 0.25 g Bromophenol Blue
40 g Sucrose
100 ml dH20
λ Hind III marker
Want a final concentration of 100 ng/µL from Invitrogen stock.
Make: 96 µL λ Hind III marker (Invitrogen stock)
320 µL TE
68 µL 6X loading buffer
20X SSC (pH 7)
175.3 g NaCl
88.25 g Na3 Citrate•2H20
1 L dH20
**pH 7.0**
4X TBS (pH 7.4)
60.55 g Tris base
4.0 g SDS
1000 ml dH20
TBS-T
1X TBS, 0.4% Tween-20
8 ml Tween-20
1992 ml 1X TBS
70
Phosphate Buffer (PBS)
4.0g NaCl
0.1g KCl
0.72g Na2HPO4
0.12g KH2PO4
In ~400ml sterile dH20
**pH 7.4**
Bring to 500ml with sterile dH20
MODULE 1
1.1: Isolation of plasmid DNA
Solution I
0.50 g D-Glucose
0.625 ml 2M Tris-Cl (pH 8)
1 ml 0.5M EDTA
Add dH20 to make total volume 50ml
**add 5 mg/ml lysozyme just before use**
Solution II
2 ml 1M NaOH
1 ml 10% SDS
7 ml dH20
**Prep fresh**
5M Potassium Acetate
29.5 ml glacial Acetic Acid
100 ml dH20
Add KOH pellets until pH=4.8
Heat-treated RNase A (100mg/ml)
Dissolve 100 mg (0.1g) of pancreatic RNase A in 1 ml 10mM Tris-Cl/15mM
NaCl. Store at -20 ºC. Immediately before use, heat RNase A in 100°C
heat block for 15 minutes and allow tubes to cool slowly to RT.
71
Salt Saturated Phenol
Tris buffered Phenol pH 6.6/7.9
8-Hydroxyquionoline added until dark yellow/orange color
Chloroform:Isoamyl Alcohol (24:1)
480 ml Chloroform
20 ml Isoamyl Alcohol
TE buffer
See general solutions section
1.4: Transformation
50 mM CaCl2
0.73 g CaCl2
100 ml dH20
** Autoclave to sterilize**
Transformation buffer
1ml 100 mM CaCl2
1 ml 100 mM Tris
1 ml 100 mM NaCl
7 ml dH20
1.6: Secondary selection of transformed bacteria
0.145 M Sterile Saline (pH ~7)
4.25 g NaCl
500 ml dH20
** Autoclave to sterilze**
72
1.8: Cracking gel
Cracking Buffer (pH 6.8)
0.788 g Tris-Cl
1.0 g SDS
0.058 g Na2EDTA•2H20
13.6 g Sucrose
0.1 g Bromophenol Blue
100 ml dH20
** pH 6.8**
1.9: Biotin labeling of DNA
3 M Sodium Acetate (pH 4.8)
24.6 g Sodium Acetate
100 ml dH20
** pH 4.8**
1.11: Southern blot
Cracking Buffer
See 1.8 above
0.5M NaOH/0.8M NaCl
20 g NaOH
46.752 g NaCl
1 L dH20
0.5M Tris/1.5M NaCl (pH 7)
250 ml 2 M Tris-base solution
87.6 g NaCl
750 ml dH20
** pH 7**
10X SSC
See general solutions for 20X SSC
73
1.13: Hybridization of Southern Blot
Prehybridization Solution (per group, prep fresh)
5 ml Formamide
2.5 ml 20X SSC
0.5 ml 100X Denhardt’s solution (doesn’t keep more than 24 hours!)
0.25 ml 1M Phosphate Buffer
200 µL Herring sperm DNA, freshly denatured
Hybridization Solution (per group, prep fresh)
4.5 ml Formamide
2.5 ml 20X SSC
0.1 ml 100X Denhardt’s solution (doesn’t keep more than 24 hours!)
0.4 ml 1M Phosphate Buffer
1.5 ml dH2O
200 µL Herring sperm DNA, freshly denatured
Biotin-labeled probe DNA
100X Denhardt’s Solution
0.2 g Ficoll
0.2 g Polyvinylpyrrolidone
0.2 g Bovine Serum Albumin (BSA)
10ml sterile dH20
**Doesn’t keep more than 24 hours**
1.14: Detection of DNA
2X SSC/0.1% (w/v) SDS (per group)
50 ml 20X SSC
450 ml dH20
0.5 g SDS
0.2X SSC/0.1% (w/v) SDS (per group)
5 ml 20X SSC
495 ml dH20
0.5 g SDS
74
0.16X SSC/0.1% (w/v) SDS (per group)
4 ml 20X SSC
496 ml dH20
0.5 g SDS
1.15: Development of Blot
Buffer 1 : Final Concentration: 0.1 M Tris-Cl
0.15 M NaCl
8.7 g NaCl
15.764 g Tris-Cl
1 L dH20
Buffer 2: 3% (w/v) BSA in Buffer 1
3g BSA per 100 ml Buffer 1
**Doesn’t keep more than 24 hours, prep fresh**
SA-AP: **Needs to be made immediately before use**
1 µL SA-AP per 1 ml Buffer 2 (approx. 10 ml needed per group)
Buffer 3
Final concentration: 0.1M Tris-Cl
0.1M NaCl
50 mM MgCl2
15.764 g Tris-Cl
5.844 g NaCl
10.15 g MgCl2
1 L dH2O
1.17: PCR
25 mM Primer Dilutions (from 500 mM stock primers)
25 µL 500mM stock
475 µL dH20
**Primers should be stored long term at 500 mM conc. and diluted to 25
mM in smaller batches**
75
1.25 mM dNTP mix (from 100 mM individual dNTP stocks)
600µL dCTP
600µL dTTP
600µL dATP
600µL dGTP
2.6 ml dH20
MODULE 2
2.2: SDS-PAGE prep and Pierce Protein Assay
4X Resolving Buffer (pH 8.9)
18.17g Tris base
10 g SDS
100 ml dH20
**pH 8.9**
Lysis Buffer
5 ml 1M Tris (pH 8.0)
3 ml 5M NaCl
1 ml Triton 100X
91 ml dH20
**add 0.0057 g DTT/10 ml Lysis Buffer fresh immediately before use**
2.3: SDS-PAGE
4X Stacking Buffer (pH 6.8)
6.055 g Tris base
0.4 g SDS
100 ml dH20
**pH to 6.8**
PAGE running buffer
10.92 g Tris base
51.36 g Glycine
3.6 g SDS
1800 ml dH20
76
6X SDS sample buffer
15 ml Glyercol
6.25 ml 2M Tris-Cl
0.186 Na2•EDTA
2 g SDS
0.1 g Bromophenol Blue
78.75 ml dH20
** 1% (v/v) β-Mercaptoethanol needs to be added fresh before use**
Coomassie Blue Stain
200 ml Methanol
50 ml Glacial Acetic Acid
1 g Coomassie Blue
250 ml dH20
Coomassie Blue Destain
200 ml Methanol
75 ml Glacial acetic Acid
725 ml dH20
2.6: SDS-PAGE and Western Blot
Western Blot Transfer Buffer (pH 8.3)
2.93 g Glycine
5.81 g Tris base
200 ml Methanol
800 ml dH20
**pH 8.3**
2.7: Blocking with primary antibody
Blocking Buffer
50 g dry milk
6.35 g Tris-Cl
1.18 g Tris base
8.77 g NaCl
1L dH20
77
TBS-T
See general solutions section.
Primary Antibody Dilution (1:5,000)
0.1% Tween-20, 1% (w/v) dry milk
50 ml 1X TBS
50 µL Tween-20
0.5 g dry milk
10 µL Anti-GST antibody
2.8: Staining with Amido Black and detection using secondary
antibody
Amido Black Stain
112.5 ml Methanol
5.0 ml Acetic Acid
0.25 g Amido Black
132.5 ml dH20
Amido Black Destain
112.5 ml Methanol
5.0 ml Acetic Acid
132.5 ml dH20
Secondary Antibody Dilution (1:10,000)
4 µL Secondary Antibody
40 ml TBS-T
MODULE 3
3.2: Overlaying plates
TBS-T
See general solutions section.
78
3.3: Identifying Positive Colonies
Blocking Buffer (0.1% (w/v) BSA)
50 g dry milk
6.35 g Tris-Cl
1.18 g Tris base
8.77 g NaCl
10 g BSA
1000 ml dH20
Primary Antibody Dilution (1:5,000)
0.1% Tween-20, 1% (w/v) dry milk
50 ml 1X TBS
50 µL Tween-20
0.5 g dry milk
10 µL Anti-GST antibody (produced in rabbit)
TBS-T
See general solutions section.
Secondary Antibody Dilution (1:10,000)
4 µL Goat anti-rabbit IgG
40 ml TBS-T
Stop Solution (pH 8.0)
1.21 g Tris-Cl
0.93 g EDTA
500 ml dH20
**pH 8.0**
MODULE 4
4.1: Making Template DNA and Performing PCR Reaction
0.9% Sterile Saline Solution
9 g NaCl
1000 ml dH20
Autoclave to sterilize
79
10% Chelex Extraction Solution
10 g Sigma Chelex 100 Resin
100 ml dH20
**prep fresh**
25 mM Primer Dilutions (from 500 mM stock primers)
25 µL 500mM stock
475 µL dH20
**Primers should be stored long term at 500 mM conc. and diluted to 25
mM in smaller batches**
1.25 mM dNTP mix (from 100 mM individual dNTP stocks)
600µL dCTP
600µL dTTP
600µL dATP
600µL dGTP
2.6 ml dH20
MODULE 5
YPD Nutrient Media
50 g YPD Nutrient Media (Difco powder)
1L dH20
**Autoclave to sterilize**
SG Buffer
9.2 g Sorbitol
1.8 g EDTA
50 ml dH20
**pH 7.4**
Phenol/Chloroform/Isoamyl Alcohol (25:24:1)
25 ml Phenol
24 ml Chloroform
1 ml Isoamyl Alcohol
**add 10mM Tris-Cl until pH=8**
80
Appendix B: Sterile Technique
It is very important in microbiology and genetics to work with pure cultures.
Unfortunately, this is difficult. The world around us is covered with
microorganisms. Microorganisms are even carried on dust particles in the air. In
order to protect sterile broth, plates, slants and pure cultures from the microbes
all around us, we must practice sterile (aseptic) technique. This simple means
that sterile surfaces or sterile media must be protected from contamination by
microbes in the air or residing on non-sterile surfaces. A simple example of the
problem is that a sterile petri plate can become contaminated with bacteria when
the lid is removed. In sterile technique, only sterile surfaces touch other sterile
surfaces and exposure to the air is kept to a minimum.
In the classroom, you often need to practice sterile technique when you inoculate
a pure culture of a microorganism into fresh medium. Sometimes this is a
transfer to a tube of liquid broth and at other times, it is a transfer to a petri platecontaining agar. While there are other circumstances that require sterile
technique, these are the most common and they will be described in more detail
on the pages that follow.
Appendix C: Spread Plate Technique
1. Dispense the appropriate volume of sample into the center of a sterile agar
plate.
2. Dip the glass spreader (aka “hockey stick”) in alcohol.
3. Pass the spreader through the flame of a Bunsen burner to burn off the
alcohol. (This sterilizes the spreader).
***IMPORTANT***
Keep the dish of alcohol behind the Bunsen burner.
Keep the alcohol dish covered when you are not using it.
Keep your hand above the spreader at all times or flaming alcohol may roll
toward your hand.
If the dish of alcohol catches on fire, cover the dish with the glass lid and it
will go out.
4. Cool the spreader by touching it to the agar where there is no sample.
5. Spread your sample over the entire surface of the agar.
6. Sterilize the spreader before putting it back on the bench.
81
Appendix D: Use of a Rainin Pipettor
Take note:
 Never rotate the volume adjustor beyond the upper or lower range of the
pipette man, as stated by the manufacturer.
 Never use the pipette man without the tip in place; this could ruin the
precision piston that measures the volume of fluid.
 Never lay down the pipette man with filled tip; fluid could run back into
the piston.
 Never let plunger snap back after withdrawing or ejecting fluid; this could
damage the piston.
 Never immerse the barrel of the pipette man in fluid.
 Never flame pipette man tips.
If you drop your pipette man, the precision piston system can be damaged;
therefore, if your pipette man is dropped, be sure to check the pipetting accuracy
has not been affected.
Recommended Volume Ranges:
Model p10: 0.5-10 µL, the number after the decimal point is in red
Model p20: 1-20 µL, the number after the decimal point is in red
Model p200: 20-200 µL, there is no decimal point
Model p1000: 200-1000 µL, the numbers after the decimal point are in black
Pipetting Directions – Method
1. Set the desired volume by holding the pipette man body in one hand and
turning the volume adjuster knob until the correct volume shows on the
digital indicator. Approach the desired volume by dialing downward
from a larger setting.
2. Press tip onto shaft by a slight twisting motion.
3. Depress the plunger to FIRST POSITIVE STOP. This part of the stroke is
the calibrated volume displayed on the digital micrometer.
4. Holding the pipette man vertically (never more than 20˚ from vertical),
immerse the tip just below the level of the liquid.
5. Allow the pushbutton to return SLOWLY to the up position. Move the tip
so that it stays slightly below the level of the liquid as you draw up.
6. Wait one to two seconds to ensure that the full volume of sample is drawn
up into the tip.
82
7. Withdraw the tip from the sample liquid.
8. To dispense the sample, place the tip end against the sidewall of the
receiving vessel and depress the plunger to the FIRST STOP. Wait one to
two seconds. Then depress the plunger to the SECOND STOP, expelling
any residual liquid in the tip.
9. With the plunger fully depressed, withdraw the pipette man from the
vessel carefully with the tip sliding along the wall of the vessel.
10. Let the plunger return slowly to the UP position. If an air bubble is
observed, re-pipette the sample.
11. Pre-rinsing the tip with the liquid being pipetted is recommended. A
significant film may be retained on the inside wall of the tip, resulting in
an error. Since the film remains relatively constant in successive
pipettings with the same tip, refilling the tip a second time and using this
quantity as the sample may obtain good reproducible results.
12. Discard the tip by depressing the tip ejector button smartly in the
appropriate waste container.
83
Appendix E: Pipette Exercises
Pipette Exercise #1
Determine and record the pipettor best suited for each of the measurements
listed below.
Add the indicated amounts to labeled microfuge tubes. Use the matrix below as
a checklist while adding solutions to each microfuge tube.
Tube A (green)
Tube B (red)
Tube C (blue)
Solution 1
10 µL
2 µL
598.6 µL
Solution 2
25µL
0.015 ml
0.200 ml
Solution 3
0.0963 ml
183 µL
201.4 µL
Determine the total volume being added to each of the tubes. To check that your
measurements are accurate, set a pipettor to the final volume and carefully
withdraw the solution from each tube. Is the tip just filled? If measurements are
inaccurate, repeat the exercise to obtain a near-perfect result.
Pipette Exercise #2
Using each of the three pipettors that are available, perform the following:
Set the pipettor to its maximum volume.
Using water at room temperature, carefully pipette the water onto a weigh boat
that you have tared (re-zeroed).
Room temperature water has a density of approximately 1 gm/ml or 1 g/L.
Therefore, you can determine the accuracy of your pipetting, e.g., 1000 L of
water will weigh 1gm. Repeat the pipetting until you feel that you are
reasonably accurate. Then record the weights of five successive pipettings.
Determine the mean and standard deviation associated with your measurements.
Complete the following conversions:
1L
=_______ml
10L =_______ml
100L =_______ml
1000L =_______ml
0.001L
0.11L
0.01ml
1L
=________ml
=________ml
=________ml
=________ml
84
APPENDIX F: PCR Reagents and Conditions
Cycling Program: GST New
94C
94C
50C
72C
5 min
1 min
1 min
1 min
Initial Elongation
94C
51C
72C
1 min
1 min
1 min
25X
72C
4C
10 min
HOLD
Final elongation
***************
5X
SAMPLES
Initial Stock
Concen.
50mM
1.25mM
25uM
25uM
5U/ul
Components
Template
DNA*
10X Buffer
MgCL2
dNTPs
Forward
Primer
Reverse Primer
Taq
Polymerase
1
2
3
4
5
3
3
colonies colonies
5
5
6
7
1-10
5
1-10
5
1-10
5
1-10
5
1-10
5
3
8
6
8
9
8
3
8
3
8
3
8
3
8
1
1
1
1
1
1
1
1
1
1
XXX
1
1
XXX
0.5
0.5
0.5
0.5
0.5
0.5
0.5
dH2O
*NOTE: Samples 1-3, 6 & 7: Use PLASMID DNA
Sample 4: Transformed Colonies
Sample 5: Non-Transformed Colonies
DESIRED FINAL VOLUME: 50µL
85
APPENDIX G: PIERCE PROTEIN ASSAY
L Alb (Stock
2mg/ml)
0
2.5
5.0
7.5
10.0
12.5
15.0
L
Sample
L dH2O
L dH2O
ml Dye
[Alb g/ml]
100
97.5
95.0
92.5
90.0
87.5
85.0
2
2
2
2
2
2
2
0
50
100
150
200
250
300
ml Dye
OD Values
g/ml in Cuvette
OD Value
Series Series
A
B







g/ml Original Solution
2
2
2
2
2
2
86
APPENDIX H: Frequently Used DNA/Protein Markers
Lambda DNA-Hind III Digest
87
APPENDIX I: GST Plasmid Map
88
APPENDIX J: Streak Plate Method
89
APPENDIX K: SDS-PAGE Assembly Instructions
90
APPENDIX L: Western Blot Assembly Instructions
91