BIOLOGY 204/205 Advanced Genetics Laboratory
TABLE OF CONTENTS
Introduction………………………………………………………………………. p. 2
MODULE 1: Recombinant DNA………………………………………………. p. 12
MODULE 2: Gene Expression…………….…………..………………………. p. 27
Appendix A: Solutions Guide……………..……………………………………. p. 39
Appendix B: Sterile Technique…………………………………………………..p. 50
Appendix C: Spread Plate Technique………………………………………... p. 50
Appendix D: Pipette Use……………………..……………………………...... p. 51
Appendix E: Pipette Exercises………………………….……………………......p. 53
Appendix F: GST Plasmid Map……………………………...……………....... p. 54
Appendix G: DNA/Protein Markers……………………………………...….. p. 55
Appendix H: Streak Plate Method …………………..……………………….... p. 56
Appendix I: PCR Reagents and Conditions for 1.17……………………….… p. 57
Appendix J: Protein Gel Setup.……………………………………………….… p. 58
Appendix K: Protein Gel Running Setup…………………………………….. p. 59
Appendix L: Pierce Protein Assay for Module 2……………………………... p.60
Appendix M: RPM to G-force Conversions………………………………….... p. 61
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BIOLOGY 204/205
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
sub-clone 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 if a cell is expressing a gene of interest,
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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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 and give a group
presentation on the results.
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 5-6 hours to finish. A few labs will
be relatively short. Students are expected to return to lab outside of the
scheduled class time, 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
6. Mini-grant presentation
Note: It is important to read over the procedures in the laboratory manual 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
occur.
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Performance in Laboratory Guidelines
1. Attendance
a. Includes arriving on time and staying until work is complete
b. Coming in willingly on “off” days when necessary
2. Working well and cooperatively with lab partner(s)
a. Simulates behavior in collaborative research groups behavior
b. Is behavior disruptive or helpful, distracting, professional?
3. Reading the lab manual in advance and arriving prepared for the day’s
methods
a. Working efficiently
b. Following through
c. Being engaged, even when not physically performing experiments
4. "Lab Citizenship"
a. Such as following the safety rules, cleaning up, labeling properly,
putting materials away, etc
5. Attitude and willingness to participate in experiments
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
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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.
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
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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 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 Proposal 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. 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.
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2. The experiment(s) will utilize molecular genetic or traditional genetic
techniques that you can do.
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.
B. Model Organisms: Your experiments must be limited to model organisms from
molecular genetics that are readily available, except mammals. Examples include:
Drosophila, bacteria, Paramecium, C. elegans, yeast, and plants.
C. Grant Proposal 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 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, single spaced, no less
than 12 point font. 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 question that will you address?
b. What are your hypotheses and specific aims?
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b. Why is this research important?
c. What has already been done in this field/topic?
d. Specifically what 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?)
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.
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. For each set of experiments 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 one
semester.) 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
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hypothesis? What are the limitations that may not allow you to have a clear
answer? This section should not exceed one page.
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 and avoid touching face and/or hair during an experiment.
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. Wear closed-toed shoes.
10. Tie back long hair.
11. Try to avoid wearing baggy, loose clothing that can interfere with your
experiment and may catch on fire.
12. Extinguish burners as soon as you finish using them.
13. All Chemical Safety and MSDS information is located in the binder on the
back of the door.
14. If you are unsure about a procedure, just ask.
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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 2 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 while 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 while 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, screw top centrifuge bottle and harvest the bacteria by
centrifuging at 5000 rpm, 4°C 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 30ml polycarbonate screw top 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. Mix gently. Let stand on
ice for 10 minutes.
6. 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.
7. Balance the tubes before centrifugation.
8. Centrifuge in the Beckman J2-21; 15,000 rpm, 4°C for 20 minutes. The
genomic DNA and bacterial debris should form a tight pellet at the
bottom of the tube.
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9. Being very careful not to disturb the pellet, divide the supernatant in half
using the p1000. Only take clear supernatant. Transfer each half to
separate 15ml glass conical tubes.
10. Add at least 500µL of heat treated RNase A to each tube. Please use all of
the RNase A provided.
11. Incubate at 37ºC for 20 minutes in Innova 4000.
12. In the chemical flow hood, add one volume of SS (salt saturated) phenol
using glass pipettes. (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, using glass pipettes.
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.
13. 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 Bench top.
14. 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) – the same amount as the
phenol you added. Vortex 1 minute and centrifuge 1 minute at 2800 rpm.
15. Transfer the upper, aqueous layer to a fresh 30ml glass tube and add 2.5
volume of cold 95% ethanol, using glass pipettes. 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!
16. Mix and allow it to precipitate on dry ice for 15 minutes.
17. Balance your tubes along with their rubber sleeves.
18. Recover the DNA by centrifuging the tube at 4°C in the Beckman J2-21 at
9500 rpm for 30 minutes.
19. Discard the supernatant into a waste container. The pellet will look like a
whitish residue on the side of the tube.
20. Resuspend the pellet with 1 ml 70% ethanol by pipetting up and down
onto the sides of the tube. Try to resuspend the entire pellet to increase
your plasmid yield. Transfer the resuspention from the first tube to the
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next tube until all pellets are resuspended and pooled together. Transfer
the solution into one sterile 1.5ml microfuge tube.
21. Microcentrifuge for 5 minutes at 14,000 rpm. Discard the ethanol; add 1ml
more of 70% ethanol to wash the pellet and vortex for 30 seconds. Spin at
14,000 rpm for 5 minutes.
22. Discard the ethanol; dry the pellet using the SpeedVac in the basement.
Give your sample to the TA/Lab tech to be properly dried for 15 – 20
minutes.
23. Dissolve the pellet in 0.3ml TE. Aliquot 100 L to each of 3
microcentrifuge tubes (properly labeled!).
24. Store at -20°C.
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.
5.
6.
7.
8.
9.
Prepare 300ml 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.
Swirl the flask and make sure all of the agarose is dissolved. If not,
microwave until it is. Remove flask with a hot mitt.
Place the running tray into the gel-casting tray. Add comb.
Cool agarose slightly; approximately 5 – 10 minutes, swirling occasionally.
Slowly pour agarose into the farthest corner from the comb in the gel
casting set up. Try to avoid bubbles! If bubbles appear remove them with
a pipette tip. Let cool until opaque (approximately 20 minutes).
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 65°C hot block.
Mix 4µL of 6X DNA sample buffer with 20µL plasmid DNA on a piece of
Parafilm.
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
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remove the comb. Removing the comb last will ensure that your wells do
not collapse.
10. Be prepared to load the gel 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 15 minutes in ethidium bromide, and
destain in water for 5 minutes.
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.
15. Leave the gel in destain to be discarded later.
16. Rinse electrophoresis unit with RO water after use so the buffer does not
dry on the electrodes.
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 cells in exponential growth phase for you to transform.
1. Divide broth culture into 2 sterile 30ml screw top 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 4°C in the Beckman J2-21 for 10 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 4°C in the Beckman J2-21 for 10 minutes
at 5000 rpm.
6. Decant the supernatant and resuspend gently 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 1.5ml microfuge tubes;
add 0.2 ml of transformation buffer to each tube. Save remaining
competent cells at 4°C.
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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. Plunge tubes into ice and let sit on ice for 5 minutes.
11. Add 0.7 ml LB to each tube and tap gently with finger.
12. Shake at 37°C for 60 minutes in Innova 4000.
**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. Let the plates dry for 5 minutes
right-side up before inverting and placing in the incubator.
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 37°C 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” LB amp 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 37°C.
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.
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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 37°C overnight for at least 24 hrs, but less than 36 hrs.
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 300ml 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 250µL of cracking buffer (Two
squares for one tube of 250µL of cracking buffer). Do this for transformed
and non-transformed cells (you should have a total of 2 microcentrifuge
tubes). Vortex tubes to mix well.
4. Incubate at 37°C 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 slowly and carefully:
Lane 1: 20µL Hind III marker (Heat in 65C hot block for 7 minutes before
loading)
Lane 2: 20µL plasmid DNA solution (10µL plasmid DNA + 4µL 6X DNA
sample buffer + 6µL 1X TAE)
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Lane 3: Transformed supernatant
Lane 4: 50µL Cracking buffer only
Lane 5: Non-transformed supernatant
Note: Load as much transformed and nontransformed supernatant as
possible (A well-formed well can hold ~50 L).
8. Run the gel for 1 hour at 100 volts.
9. Stain with ethidium bromide, destain, and photograph. Look for genomic
DNA, plasmid DNA and RNA.
1.9 Labeling DNA with Biotin
-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 is to be used only wearing gloves under the hood. Dispose of all
pipets and liquid waste containing chloroform in appropriate disposals.
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):
dH2O
128µL
dNTP mix
28µL
1X DNase I Buffer
19.9µL
DNase I
0.1µL
Plasmid DNA
4µL
DNA Polymerase I
20µL
3. Mix well and centrifuge for 5 seconds at 14,000 rpm.
4. Allocate 50µL into 4 tubes.
5. Incubate at 15°C for 2 hours in thermocycler.
6. Add 5µL Stop Buffer to each tube and mix.
7. 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
18
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. Collect top liquid layer and transfer to a clean tube. Discard remaining
liquid into waste container.
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 5 minutes 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 on lab bench 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.
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
non-transformed 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. Run cracking gel (same as Day 1.8, except add 20ul of plasmid mixed with
4ul 6X DNA sample buffer to the plasmid lane). Do not forget control
lane!
19
2. Stain with ethidium bromide, briefly destain, and examine the gel.
3. Make sure to destain the gel for approximately 5 minutes before
denaturing.
4. 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 Spot.
a. Take your Biotin labeled probe out of the freezer and let
thaw on ice.
b. Cut a piece of nitrocellulose approximately 1 cm x 1 cm.
Make sure to cut your test spot in a unique way so that you
can identify it later. For example you can cut one or two
small notches on the side of the square or cut off a corner.
c. Hydrate the test spot in dH2O for 3 minutes.
d. Soak the nitrocellulose in 10X SSC until the probe is thawed.
e. Remove the nitrocellulose from the SSC and place on a small
Kimwipe.
f. Add 2 µL of probe to the center of the square of
nitrocellulose.
g. Let dry on Kimwipe, then wrap in plastic wrap and store in
the freezer until 1.12.
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 pieces 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**
20
-Place three more pieces 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 placed on top of the filter paper (All filter paper and paper
towel should be cut to the size of the gel).
-Wrap the whole set up in plastic wrap to provide stability to the
stack and minimize evaporation.
-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. Disassemble the Southern blot and rinse the nitrocellulose in 5X SSC for 2
minutes.
2. Dry on large Kimwipe.
3. Bake nitrocellulose blot on a kimwipe and test spot wrapped in plastic in
vacuum oven at 80°C for 2 hours.
4. Carefully place blot into hybridization bag and seal on all four sides.
5. Store blot and test spot in freezer.
1.13 Hybridization of the Southern Blot
1. Carefully unwrap your test spot.
2. Place your uniquely cut test spot in a small plastic tub containing all of the
test spots from the class. Your TA or lab tech will hydrate these in 2X SSC
and then place them in prehybridization solution and return them to you
tomorrow for 1.14.
3. Cut a corner of your hybridization bag. Using a stereological pipette, add
50ml 2XSSC to the bag to hydrate your blot. Reseal the corner of the bag
using the food sealer. The blot should be uniformly hydrated after several
minutes.
4. While the blot hydrates, denature 200L of Herring sperm DNA
(2mg/mL) in 100°C hot block for 10 minutes followed by plunging into ice
water.
5. For prehybridization of the nitrocellulose blot, add the 200µL of freshly
denatured Herring sperm to the prehybridization solution and mix.
6. Cut a corner of your bag and pour in the prehybridization solution using a
stereological pipette. Reseal the corner making sure to push all of the air
bubbles out of the bag. If there are still more air bubbles in the bag after
21
you have resealed the corner, set the bag upright and push all the bubbles
to the top of the bag. Reseal the bag across the top to trap the air bubbles
away from the blot.
7. Incubate at 42°C while rocking for 2 hours. The volume of
prehybridization solution used should be 20 to 100L per cm2 of the blot.
8. For hybridization, heat-denature 5L of the probe made on 1.9 and 200 L
of Herring sperm DNA by placing in 100°C hot block for 10 minutes.
While in the heat block, wrap tops of tubes in Parafilm to preserve the
label in the next step.
9. Plunge into ethanol ice slurry for fast chilling, making sure not to erase all
labels written in marker. Just before use, add to the hybridization solution.
10. Remove the prehybridization solution from the bag by cutting a corner
and pouring off. Add the hybridization solution to the bag (20-100 L per
cm2) and reseal using the same techniques described in step 6. The blot
should be hybridized at 42°C overnight while rocking 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 blot & test spot with 100ml of 2X SSC/0.1% (w/v) SDS at room
temperature for 3 minutes. Decant the SSC and repeat.
2. Wash the blot & test spot 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 blot & test spot in 100ml of 0.16X SSC/0.1% (w/v) SDS at 50°C
for 15 minutes. Decant the SSC and repeat.
4. Rinse the blot & test spot 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.
22
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 blot and small test square in Buffer 1 at room temperature for 1
minute with sufficient buffer to cover. Decant Buffer 1 into the sink.
2. Incubate blot and test spot in Buffer 2 in a plastic container for 1 hour at
65°C, rocking, with sufficient buffer to cover. Then decant off.
3. Wash the blots in freshly made strep-avidin alkaline phosphatase (SA-AP)
conjugate for 25 minutes at room temperature. 1µL SA-AP per 1mL
Buffer 2.
(Add only enough SA-AP conjugate to cover the blots (~10ml). Use gentle
agitation and occasionally pipette SA-AP over the blots.)
4. Decant and save the SA-AP in a 15ml tube. Save for step #6.
Wash the blot and test spot in Buffer 1 using 20 to 40-fold greater volume
than employed in step 3. Gently agitate blot for 15 minutes in Buffer 1.
(if you used 10 ml diluted SA-AP conjugate in step 3, wash with at least 200400ml Buffer 1.) Decant Buffer 1 into the sink.
5. Wash the blots for 10 minutes in Buffer 3, rocking. Decant Buffer 3 into
sink.
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. Add 9 ml of NBT-BCIP solution to the blots. Allow the blots to develop
for 15 minutes to 1 hour. (Do steps 6 and 7 at the same time and monitor
the rate of color development. The tube of saved SA-AP acts as a positive
control.) Agitate the Tupperware.
8. DNA bands will be most evident on only one side of the blot (check your
notch for correct orientation). Check your blot every 2 minutes to ensure
that over-development does not occur.
9. Once bands have developed, decant the NBT-BCIP solution in the
appropriate waste container and wash the blot in TE. This will terminate
the color development reaction. The TE can then be decanted into the
sink.
10. Let the blots dry on a large Kimwipe. Then wrap in plastic wrap and
label. The lab tech will photograph and distribute the blots for your
notebooks.
23
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.
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.
MSPILGYWKIKGLVQPTRLLLEYLEEKYEEHLYERDEGDKWRNKKFELGLEFP
NLPYYIDGDVKLTQSMAIIRYIADKHNMLGGCPKERAEISMLEGAVLDIRYGV
SRIAYSKDFETLKVDFLSKLPEMLKMFEDRLCHKTYLNGDHVTHPDFMLYDA
LDVVLYMDPMCLDAFPKLVCFKKRIEAIPQIDKYLKSSKYIAWPLQGWQATFG
GGDHPPKSDLVPRGSPGIHRD
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 I. 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 color and tube number!
24
2. For Sample 4, pick up three individual colonies from your transformed
plate with a sterile toothpick and place into a 1.5 ml microfuge tube filled
with 50 µL of sterile dH2O. Take 3µL of this bacterial solution and use as
your “template DNA”. Do the same for Sample 5 using non-transformed
bacteria.
3. Before mixing the reactants, 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 template DNA that you add might differ from tube
to tube.
4. 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.
5. Add all reactants, except the Taq, while the tubes are on ice.
6. Lastly add the Taq polymerase.
7. Once all reactants 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.), 0.5
ml tubes are thin-walled and can crack if microfuged for too long.
8. Keep the tubes on ice until the entire class is ready to load the
thermocycler.
9. 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. Note: The 2% agarose solution will solidify
quickly! Pour gel while still relatively hot.
3. Remove 20µL of PCR product from each tube; add to 4 µL of 6X sample
buffer. Store the remaining PCR product at 4°C.
4. Once your gel is set, remove the comb and place the gel in the running
box. Cover the gel with 1X TAE buffer.
5. Load your DNA samples and 10µL of 100bp ladder into the gel.
6. Run gel at 100 volts for 1 hour.
25
7. Once the electrophoresis is complete, stain your gel for ~15 minutes in
ethidium bromide. WEAR GLOVES! Ethidium bromide is a mutagen
and carcinogen.
8. Destain, examine, and photograph 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
26
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 SulfatePolyacrylamide 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 SDS-PAGE prep - Pour Resolving Gel______________________
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.
1.
2.
3.
4.
Obtain one 1.5mm spacer glass plate and one short glass plate; wipe
with methanol and a Kimwipe until you hear a “squeaky” noise.
Handle glass plates at edges. Wear gloves!
Assemble the gel casting apparatus (See diagram in Appendix J).
Assemble on a flat surface and then clamp shut.
Insert the 1.5mm comb and use a Sharpie to draw a line across the
glass 0.7 cm below the comb. Once the line is drawn, remove the
comb.
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. Make sure to remove the water before pouring the gel. Use
a Kimwipe to remove water droplets from between the plates of glass.
Before you pour your gel, have the TA or lab tech check your apparatus!
27
5. Prepare a 12% 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
6. Gently swirl the solutions to mix WELL.
7. 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.
8. Overlay the acrylamide with dH2O. Do this by gently adding the dH2O
with a glass Pasteur pipette. You will be able to see a distinct line between
the dH2O and the resolving gel solution.
9. 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).
10. Wrap the gel/casting apparatus in a damp paper towel and then plastic
wrap. Label appropriately. Store in the cold room.
2.2 Preparation of bacteria for SDS-PAGE (Done for you)_________
1. Using a sterile toothpick, select one colony from a plate of freshly
transformed BL21+GST cells and add to 10ml of LB-amp broth. Shake
tube at 37C ON.
2. Add one control (non-transformed) BL21 colony to 10ml LB broth. Shake
tube at 37C ON.
2.3 SDS-PAGE preparation and Pierce Protein Assay
Four hours before class …inoculate bacterial cultures (Done for you):
1. Inoculate 2 tubes of 10ml of LB amp broth with 500L of transformed ON
bacterial culture each.
2. Inoculate 2 tubes of 10ml of LB broth with 500L control (nontransformed) ON bacterial culture each.
3. Shake at 37C for 1 hour.
4. To ONE of the LB amp cultures and ONE of the LB cultures, add 300L of
100mM IPTG each. Label tubes to indicate the addition of IPTG.
5. Shake at 37C for an additional 3 hours.
28
Prepare the proteins for the gel and protein assay:
1. Obtain 2 ml of each type of culture (Transformed, Transformed + IPTG,
Untransformed, and Untransformed + IPTG). Split each culture into two
1.5 ml microcentrifuge tubes. You will have a total of 8 tubes (2 x
Transformed -IPTG, 2 x Transformed +IPTG, 2 x Untransformed-IPTG, 2 x
Untransformed +IPTG).
2. Spin all tubes in the microcentrifuge at 14,000 rmp for 1 minute.
3. Decant the supernatant from each tube.
4. Resuspend one of each kind of pellet (i.e. transformed, transformed plus
IPTG , -control, and +control) in 100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 4 tubes.
6. Sonicate each sample in the cold room, 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. Keep protein on ice for protein assay.
9. After setting up protein assay, freeze the remaining samples at -20C.
Label appropriately!
Pierce Protein Assay (See directions in Appendix L)
Use the chart provided to develop a standard curve using BSA standards and to
determine protein concentration. Perform the following procedure in duplicate!
(You will run two sets of protein assays and average the OD values to calculate
the protein concentrations.)
1.
Dilute protein sample: Make 10X and 20X dilutions for each protein
sample. For example, to make a 20X dilution, add 5L of your sample to
95L of sterile dH2O. For a 10X dilution add 10L of your sample to 90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. Mix the dye in glass jar provided.
3. Add BSA and dH2O according to the directions in Appendix L.
4. Add 2ml of the dye to each one of your samples and standards.
29
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.
12. Establish a standard curve using the OD values obtained with your BSA
standards: graph Concentration (x-axis) vs. OD (y-axis) on graph paper.
Using this graph, calculate the protein concentrations in your four
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.
2.4 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
30
4X Stacking Buffer pH 6.8
30% Acrylamide stock
TEMED
2.50 ml
1.30 ml
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 ethanol 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.
9. Once the gels are set, remove them from the casting stand and assemble in
the gel box (See Appendex K). Note: Do not remove the comb yet.
10. Add 1X PAGE Running buffer to the upper chamber. The buffer level
should be half way between the top of the big and small glass plates.
11. Add 1X PAGE Running buffer to the lower chamber until the appropriate
level for the number of gels in the box is reached.
12. Carefully remove the comb.
Gel Set Up
Your polyacrylamide gel will have 10 lanes. The first five lanes will contain a
protein marker and your four protein samples. The last five lanes will contain
the same set of protein samples in the same order. 20µg of each protein sample
should be added per lane.
1. Calculate the volumes of sample, sample buffer, and water needed for
each tube (make 60µl total so that even if some evaporates during boiling,
there will still be 40µl left; 20µl per lane):
-Each tube needs 60µg of protein total. Using the concentrations
you calculated off of your standard curve, calculate how many
microliters equals 60µg.
-Each tube needs 1X SDS sample buffer. You are given 6X SDS
sample buffer. Calculate how much 6X SDS sample buffer is
needed so that the final concentration is 1X.
31
2.
3.
4.
5.
6.
7.
-Each tube needs a total volume of 60µl. Figure out how much
water should be added to each so that the total volume is 60µl.
Add the calculated volumes of water, protein, and 6X SDS sample buffer
to a 1.5ml tube, in that order. Make sure to mix sample buffer and
protein samples before using! Place the remaining protein samples in the
freezer.
Boil samples for 5 minutes right before you are ready to load. After
boiling, keep samples on ice while loading gel.
Using gel loading tips, load your samples into the gel in the following
order (load 20µl per lane):
Lane 1, 6: Prestained protein marker
Lanes 2, 7: Transformed
Lanes 3, 8: Transformed plus IPTG
Lanes 4, 9: Control (non- transformed)
Lanes 5, 10: Control (non- transformed) plus IPTG
Run the gel at 50mA for approximately 1 hour; until the dye hits the
bottom edge of the gel.
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.
Put the gel into a plastic container and cover with Coomassie Blue Stain.
Rock gently overnight.
2.5 Dry Gel (Done for you)
1. Transfer gel to destain. Wash and discard destain in correct waste
container.
2. Rinse gel with destain again and decant destain into correct waste
container.
3. Submerge the gel in more destain and rock gently for 1 hour or until
background of gel is transparent.
4. Discard destain in appropriate waste container.
5. Rinse gel in dH2O. Discard into appropriate waste container.
6. Submerge your gel in gel drying buffer and place back on the shaker ON.
7. Photograph and appropriately dry gels using cellophane.
8. Make sure to observe and record gel appropriately.
32
2.6 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.
2.
3.
4.
Obtain one 1.5mm spacer glass plate and one short glass plate; wipe
with methanol and a Kimwipe until you hear a “squeaky” noise.
Handle glass plates at edges. Wear gloves!
Assemble the gel casting apparatus (See diagram in Appendix J).
Assemble on a flat surface and then clamp shut.
Insert the 1.5mm comb and use a Sharpie to draw a line across the
glass 0.7 cm below the comb. Once the line is drawn, remove the
comb.
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. Make sure to remove the water before pouring the gel. Use
a Kimwipe to remove water droplets from between the plates of glass.
Before you pour your gel, have the TA or lab tech check your apparatus!
5. Prepare a 12% 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
6. Gently swirl the solutions to mix WELL.
7. 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
33
until it is just barely above your Sharpie line. Try to avoid making
bubbles.
8. Overlay the acrylamide with dH2O. Do this by gently adding the dH2O
with a glass Pasteur pipette. You will be able to see a distinct line between
the dH2O and the resolving gel solution.
9. 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).
10. Wrap the gel/casting apparatus in a damp paper towel and then plastic
wrap. Label appropriately. Store in the cold room.
2.7 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 (makes enough for 2 gels).
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, pour the dH2O off the resolving
gel and dry with a Kimwipe.
4. Add the ammonium persulfate to your tube. Mix gently.
34
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 ethanol 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 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 30 minutes for the gel to polymerize. Thaw your protein samples
on ice while waiting.
8. Once the gels are set, remove them from the casting stand and assemble in
the gel box (See Appendex K). Note: Do not remove the comb yet.
9. Add 1X PAGE Running buffer to the upper chamber. The buffer level
should be half way between the top of the big and small glass plate.
10. Add 1X PAGE Running buffer to the lower chamber until the appropriate
level for the number of gels in the box is reached.
11. Carefully remove the comb.
Gel Set Up:
1. Calculate the volumes of sample, 6X SDS sample buffer (mix before
using), 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. Boil samples for 5 minutes right before you are ready to load. After
boiling, keep samples on ice while loading gel.
4. Using gel loading tips, load your samples into the gel in the following
order (load 20µl per lane):
Lane 1, 6: Prestained protein marker
Lanes 2, 7: Transformed
Lanes 3, 8: Transformed plus IPTG
Lanes 4, 9: Control (non- transformed)
Lanes 5, 10: Control (non- transformed) plus IPTG
5. 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.
35
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!
5. Soak the blotting paper in Western blot transfer buffer.
6. Rinse the nitrocellulose membrane with dH2O, and 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 a
large Kimwipe. 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.
36
2.8 Primary Antibody Blocking (day before 2.9)
1. Cut the nitrocellulose along the edge of the middle prestained marker so
that both halves of the nitrocellulose contain the visible protein 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 in 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 1X TBS for five minutes, rocking. Decant into
sink. Repeat for a total of 3 five minute washings.
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.
2.9 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.
-NBT/BCIP is highly toxic. WEAR GLOVES when handling and dispose of all
liquid waste containing NBT/BCIP in the appropriate waste container.
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 destain into the proper waste container.
5. Place the blot on Kimwipe 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. Wash the blot with TBS-T for 5 minutes, rocking. Decant the TBS-T into
the sink. Repeat 2 more times.
3. Add the secondary antibody (Goat anti-rabbit IgG alkaline phosphatase
conjugated diluted 1:10000 in TBS-T).
37
4. Rock the blot at room temperature for 1 hour. Save a small volume
(500L) of the secondary antibody in a 1.5 ml tube to use as a control, keep
on ice.
5. Decant the rest of 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 post-antibody 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. (Should occur within 1 to 10 minutes).
10. Once color has developed, decant the NBT-BCIP solution into its
appropriate waste container. Rinse the nitrocellulose with TBS-T. Decant
into the sink.
11. Air dry the nitrocellulose on a Kimwipe. 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.
38
Appendix A: Solutions Guide
GENERAL SOLUTIONS
Ampicillin Stock
100 mg/ml stock solution: for example, 0.5g ampicillin sodium salt into 5
ml dH2O. Filter sterilize, and store at -20°C.
LB amp
100 µg/ml final concentration: 1:1,000 dilution of ampicillin stock into LB
broth.
When making LB amp plates, add 1 ml ampicillin stock (100mg/ml) into
1L LB Agar broth. *NOTE: Ampicillin is heat-sensitive, so LB agar 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 agar
broth cool in there for an hour is a good way to ensure the LB agar doesn’t
solidify.
When making LB amp broth, add 1ul of ampicillin stock (100mg/ml) for
each 1ml of LB broth.
TE buffer
Need (final conc.): 10mM Tris-Cl (pH 7.5), 1mM EDTA (pH 8.0)
Make from liquid stocks of Tris-Cl and EDTA
5ml 2M Tris-Cl (pH 7.5)
2ml 0.5M EDTA (pH 8)
993ml dH2O
2M Tris-Cl (pH 8.0)
177.6g Tris-Cl
10.6g Tris-base
In ~950mL sterile dH2O
**pH 8.0**
Bring up to 1L with sterile dH2O
39
0.5M EDTA (pH 8)
18.6g EDTA disodium salt (FW= 372.2)
In ~75ml sterile dH2O
Heat in microwave to dissolve salt
***bring pH to 8.0***
Bring up to 100mL with sterile dH2O
50X TAE stock (pH 8.5)
242g Tris Base (FW= 121.14)
In ~700ml sterile dH2O
Carefully add 57.1mL Glacial Acetic Acid
100mL 0.5M EDTA (pH 8.0)
Bring up to 1L with sterile dH2O
pH 8.5, but no adjustment needed
*Dilute 50X TAE stock 1:10 for a 5X stock*
6X DNA sample buffer
0.25 g Bromophenol Blue
40 g Sucrose
100 ml dH2O
λ Hind III marker
100 µL λ Hind III Digest (NE Bio Labs stock)
150 µL TE
50 µL 6X DNA sample buffer
20X SSC (pH 7)
175.3 g NaCl
88.25 g Na3 Citrate•2H2O
1 L dH2O
**pH 7.0**
2X SSC
10ml 20X SSC
90ml dH2O
4X TBS (pH 7.4)
60.55 g Tris base
4.0 g SDS
1000 ml dH2O
40
TBS-T
1X TBS, 0.1% Tween-20
2 ml Tween-20 (use large orifice tips to pick up Tween)
1998 ml 1X TBS
Phosphate Buffer (PBS)
4.0g NaCl
0.1g KCl
0.72g Na2HPO4
0.12g KH2PO4
In ~400ml sterile dH2O
**pH 7.4**
Bring to 500ml with sterile dH2O
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 dH2O 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 dH2O
**Prep fresh**
5M Potassium Acetate
29.5 ml glacial Acetic Acid
100 ml dH2O
Add KOH pellets until pH=4.8
Store in refrigerator
41
Heat-treated RNase A (100mg/ml)
Dissolve 600 mg (0.6g) of pancreatic RNase A (Sigma R-4875) in 6 ml
10mM Tris-Cl/15mM NaCl. Divide into 6 1.5ml microfuge tubes and heat
in 100°C heat block for 15 minutes. Allow tubes to cool slowly to room
temperature. Store at -20 °C.
10mM Tris-Cl/15mM NaCl
0.1576 g Tris-Cl
0.0876 g NaCl
100ml dH2O
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 dH2O
** Make fresh**
Transformation buffer
1ml 100 mM CaCl2
1 ml 100 mM Tris
1 ml 100 mM NaCl
7 ml dH2O
**Store at 4°C**
42
1.6: Secondary selection of transformed bacteria
0.145 M Sterile Saline (pH ~7)
4.25 g NaCl
500 ml dH2O
** Autoclave to sterilize**
1.8: Cracking gel
Cracking Buffer (pH 6.8)
0.788 g Tris-Cl
1.0 g SDS
0.058 g Na2EDTA•2H2O
13.6 g Sucrose
0.1 g Bromophenol Blue
100 ml dH2O
** pH 6.8**
1.9: Biotin labeling of DNA
3 M Sodium Acetate (pH 4.8)
24.6 g Sodium Acetate
100 ml dH2O
** 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 dH2O
0.5M Tris/1.5M NaCl (pH 7)
250 ml 2 M Tris-base solution
87.6 g NaCl
750 ml dH2O
43
** pH 7**
10X SSC
See general solutions for 20X SSC
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 (2mg/ml, made fresh), 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 (2mg/ml), 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 dH2O
**Doesn’t keep more than 24 hours**
1.14: Detection of DNA
2X SSC/0.1% (w/v) SDS
50 ml 20X SSC
450 ml dH2O
0.5 g SDS
44
0.2X SSC/0.1% (w/v) SDS
5 ml 20X SSC
495 ml dH2O
0.5 g SDS
0.16X SSC/0.1% (w/v) SDS
4 ml 20X SSC
496 ml dH2O
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 dH2O
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
45
1.17: PCR
20 µM Primer Dilutions (from 500 µM stock primers)
5 µL 500µM stock
120µL dH2O
**Primers should be stored long term at 500 µM conc. and diluted to 20
µM in smaller batches**
10 mM dNTP mix (from 100 mM individual dNTP stocks)
500µL dCTP
500µL dTTP
500µL dATP
500µL dGTP
3 ml dH2O
*1 µL of 10mM dNTP mix per 50µL reaction*
1.25mM dNTP mix
262.5µL dH2O
37.5µL 10mM dNTP mix
*8 µL of 1.25mM dNTP mix per 50 µL reaction*
MODULE 2
2.2: SDS-PAGE prep and Pierce Protein Assay
100mM IPTG
0.02383g IPTG
1 ml dH2O
4X Resolving Buffer (pH 8.9)
18.17g Tris base
10 g SDS (measure in hood)
100 ml dH2O
**pH 8.9**
46
Lysis Buffer
5 ml 1M Tris (pH 8.0)
3 ml 5M NaCl
1 ml Triton 100X
91 ml dH2O
**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 dH2O
**pH to 6.8**
5X Running Buffer
7.55 g Tris Base
36 g Glycine
Bring solution to 500 ml volume and stir
pH should be 8.3 but don’t adjust
Add 2.5 g SDS
**Store at 4°C**
Dilute to 1X before using
6X SDS sample buffer
7 ml 4X Tris-Cl/SDS, pH 6.8
3.8 ml glycerol (30% final)
1 g SDS (10% final) (weigh in hood)
2 mg bromophenol blue
Add dH2O to 10 ml volume
*Store in .5 ml aliquots at -20*
Add 50µL β-ME (5%) to each 1ml tube before use
4X Tris-Cl/SDS, pH 6.8 (0.5 M Tris-Cl with 0.4% SDS)
6.05g Tris Base in 40 ml dH2O
Adjust pH to 6.8 with 1M HCl
Add dH2O to 100 ml total
Add 0.4 g SDS (weigh in hood)
*Store for up to 1 month*
47
Coomassie Blue Stain
200 ml Methanol
50 ml Glacial Acetic Acid
1 g Coomassie Blue
250 ml dH2O
Coomassie Blue Destain
200 ml Methanol
75 ml Glacial acetic Acid
725 ml dH2O
2.4: Dry Gel
Gel Drying Buffer
50 ml Glycerol (10%)
35 ml Acetic Acid (7%)
125 ml Methanol (25%)
290 ml dH2O (58%)
*Glycerol is added first in a large graduated cylinder, mix well with water.
Add the acid last.*
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 dH2O
**pH 8.3**
2.7: Blocking with primary antibody
Blocking Buffer
5 g dry milk
.635 g Tris-Cl
.118 g Tris base
.877 g NaCl
100ml dH2O
48
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 dH2O
Amido Black Destain
112.5 ml Methanol
5.0 ml Acetic Acid
132.5 ml dH2O
Secondary Antibody Dilution (1:10,000)
4 µL Anti-Rabbit IgG
40 ml TBS-T
49
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 simply 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.
50
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.
51
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.
52
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 the p1000 and the p200 pipettors, 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
53
APPENDIX F: GST Plasmid Map
54
APPENDIX G: Frequently Used DNA/Protein Markers
Lambda DNA-Hind III Digest
100 bp DNA Ladder
Prestained Protein Marker
1Kb DNA Ladder
55
APPENDIX H: Streak Plate Method
1
2
3
4
5
Flame loop in between each step
except between 4 and 5.
Do not flame loop between steps 4
and 5.
56
APPENDIX I: PCR Reagents and Conditions for 1.17
Cycling Program: GST
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
Concentration
25mM
1.25mM
20µM
20µM
5U/µl
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
57
APPENDIX J: Protein Gel Setup
58
APPENDIX K: Protein Gel Running Setup
59
APPENDIX L: PIERCE PROTEIN ASSAY for Module 2
Label
S1
S2
S3
S4
S5
S6
S7
Label
L Alb (Stock
2mg/ml)
0
2.5
5.0
7.5
10.0
12.5
15.0
L
Sample
L
dH2O
100
97.5
95.0
92.5
90.0
87.5
85.0
L
dH2O
ml
Dye
ml
Dye
2
2
2
2
2
2
2
OD Value
Series
Series
A
B
[Alb
g/ml]
0
50
100
150
200
250
300
OD Values
g/ml in
Cuvette
g/ml
Original
Solution
2
2
2
2
2
2
2
2
Label
Average g/ml
Original Solution
of each treatment
g /l
Original
solution
l of protein
that equates to
60g total
Transformed
Transformed + IPTG
Control
Control +IPTG
60
APPENDIX M: RPM to G-Force Conversions
Equipment
Beckman J2-21 w/JA 14 rotor (250mL tubes)
Beckman J2-21 w/JA 17 rotor (30 mL tubes)
Beckman J2-21 w/JA 17 rotor (30 mL tubes)
Beckman J2-21 w/JA 17 rotor (30 mL tubes)
IEC Centra 7 Desktop w/ 15 mL tubes
IEC Centra 7 Desktop w/ 50 mL tubes
Eppendorf Centrifuge 5702 w/ 15 mL tubes
Eppendorf Centrifuge 5702 w/ 50 mL tubes
Eppendorf Centrifuge 5415 C w/ 1.5 mL tubes
Sorvall Legend Centrifuge w/ 1.5 mL tubes
RPM
G-Force (RCF)
5,000
3,840
5,000
3,440
9,500
12,400
15,000
31,000
2,800
1,098
2,800
1,098
2,800
1,120
2,800
1,180
14,000
15,980
14,000
18,800
61