Manual For Chemical Biology Laboratory
Manual For Chemical Biology Laboratory
Manual For Chemical Biology Laboratory
Chemistry 427b
Spring 2004
page 1
1. Course Description (return to Contents)
The goal of Chemical Biology Laboratory is to involve undergraduates enrolled in
Chemical Biology I in the challenge and excitement of independent discovery at an early
stage of their Yale experience, long before such research opportunities would usually be
available to them. Each student will have an individual project that is a sub-project of a
major funded investigation currently underway in graduate research laboratories at Yale.
Since the projects in this course represent novel research, they differ from those in a
traditional laboratory course which tend to have pre-determined outcomes. Students in
Chemical Biology I will share the excitement—and perhaps the frustration—of hands-on
experience with original research.
As students progress through the semester, they will gain experience performing a variety
of indispensable laboratory techniques while they gain exposure to research methods. By
the end of the semester, students will be expected to analyze their results and propose
logically related future experiments.
The laboratory classroom is SCL 168, with sub-groups meeting from 1 – 5:00 p.m. on
either a Monday-Wednesday or a Tuesday-Thursday schedule. Sub-groups will meet
from 1 – 1:30 p.m. in SCL 3 with the TA and/or Dr. Allen for pre-lab discussions on
MW or TTh (according to their scheduled laboratory sessions). A 50-minute Friday
lecture will be held from 2 – 2:50 p.m. in SCL 3. All Chemical Biology Laboratory
students are encouraged (though not required) to attend the Schepartz laboratory group
seminars, which are held at 3 p.m. in SCL 201 (the Faculty Lounge).
Note: Each sub-group will have distinct schedule guidelines for experimental progress.
Because of the inherently open-ended, results-driven nature of research, it is impossible
to provide an exact script of the semester! This is part of the excitement of research, but it
also requires students to maintain focus and look to their TA for direction as necessary.
Other than how experimental time is organized, the rest of the schedule is identical for all
sub-groups. Every student is responsible for attending all scheduled laboratory and
lecture periods. If an illness, injury, or family emergency prevents you from fulfilling this
obligation, please email the instructor and your TA before the absence.
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Below is the general schedule that applies to all sub-groups (see next page). Click the
links that follow to view the specific schedule for your particular sub-group.
Sub-group 1: The β-peptide project: What is the effect of salt bridge structure on
14-helix stability?
Sub-group 3: The phage display project: Can we identify miniature proteins that
bind human MDM2 with high affinity?
page 3
Chemical Biology Laboratory Weekly Schedule
page 4
3. Course Materials (return to Contents)
Lab text:
This lab manual serves as the guiding text for the course. Each sub-group description will
contain references to relevant techniques and background journal articles from the
literature. There are also several recommended textbooks to provide background
information for many of the techniques and biological processes with which you will
become acquainted during this course.
Recommended Textbooks:
In addition to the textbooks listed above, each sub-group has a list of relevant journal
articles for background and supplementary reading. Most of these are available online
for free to Yale IP addresses.
Click on the link below for the list of literature specific to your sub-group project:
*Section 13 below contains links to information about the protocols and laboratory
techniques you will be using.
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Laboratory notebook: Available at the Yale bookstore, or you may use a lab notebook
begun in a previous course. The notebook cannot be spiral or loose-leaf, and must be
capable of making carbon copies to hand to your TA.
Calculator: You will need a reliable scientific calculator for calculation of yields,
concentrations, molarity conversions, etc. To prevent loss, please label it with your name.
Lab safety glasses or goggles: Eyewear will be provided. Students who wear glasses
should wear goggles or safety glasses over them.
Lab coat: This is suggested but optional, and you may instead opt to wear durable,
inexpensive clothing that you don’t mind staining.
Students will be graded on their understanding and performance of the techniques that are
entailed in the class, the quality of their experimental design, laboratory notebooks,
experimental data, and their ability to draw conclusions from results and place them in a
scientific context. There will be four quizzes on material presented in the Friday lectures,
a written proposal for future experiments, and a final report in the format of a short
Journal of the American Chemical Society article for the final exam.
Breakdown:
20% 4 quizzes on lecture material
10% Pre-lab assignments
40% Laboratory notebooks
10% Proposal for future experiments
20% Final report (format of short JACS article)
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Academic Honesty
Honor Code
As an enrollee in this chemical biology laboratory course, I agree to work independently unless I
am specifically instructed to work with a partner. I will not copy another student’s work on any of
the assignments or quizzes. I will not allow another student to copy my work on any of the
assignments or quizzes. In my laboratory notebook, I will record all data with honesty and submit
only my own work, unless I have clearly credited another student for contributing data in
collaboration.
Faculty
Unless otherwise informed by your TA, you will meet at the beginning of each laboratory
session in SCL 3 for a brief discussion about the day’s activities. The laboratory
classroom is SCL 168.
The lab doors are LOCKED outside of scheduled lab periods. With your TA’s
permission, you are welcome to additional lab time during another scheduled lab period,
or at another time that you arrange with your TA. There may be occasions when your
experiments require attention outside of your scheduled lab period, and in that eventuality
you will be invited (but not required) to participate.
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6. Lab Safety and Waste Management (return to Contents)
Safety
Your safety is of primary importance. You should always arrive prepared to work
effectively, with an experimental plan for the day and basic knowledge of the
instruments, materials, and techniques you will be using. In addition to thorough
preparation, know the cardinal rules of lab safety listed below. Safety Rules are also
posted at each TA blackboard and at the bulletin board by the organic lab (145 SCL).
Safe Housekeeping Rules are posted at each chemical ventilation hood.
Although you will not be working with any live pathogens or infectious materials, you
should familiarize yourself with the basic guidelines for biological safety and waste
management (http://www.yale.edu/oehs/LabIssues/Bio/bioreqmain.htm; see Section X,
Biological Waste Disposal). Additional information on biological safety can be found at
http://www.cdc.gov/od/ohs/biosfty/biosfty.htm.
Finally, your lab TA will provide information about lab safety and waste management for
specific experiments and techniques. If you don’t know, ASK.
You are expected to know and follow the safety rules listed below. In addition, Chemical
Biology Laboratory students are responsible for following the guidelines for safe
housekeeping, also listed below. This course gives you more independence than other
undergraduate laboratories, and maintaining a clean and organized work environment is
important for conducting experiments safely and efficiently.
Safety Rules:
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Notify your TA/staff immediately in case of accident, injury, fire, leak, or
chemical/water spill.
Wear safety goggles or safety glasses with side shields at all times in the lab.
Know the location and operation of the shower and eyewash in your lab.
Know the location of the fire extinguisher and all exits in the lab.
Use chemical reagents under the chemical ventilation hoods, and take care to
recap the bottles after use.
LABEL your tubes, beakers, buffers, and other solutions to avoid mix-ups.
Dispose of solid and liquid waste products in the correct waste container. If you
are not sure how to deal with waste, ask your TA.
Shut off gas, water, steam and electrical devices before leaving the lab.
Never perform any unauthorized experiments. Always ask your TA before you
try something novel.
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If you break it or use it up, replace it or notify your TA/staff.
Do your part to keep the hoods, benches, reagent containers, and equipment
clean by wiping up spills immediately.
Avoid risk of flooding by keeping all trash and debris out of the sinks.
Clean your glassware and bench space at the end of the lab day.
Waste
Proper waste management is an important part of safe laboratory practice. Guidelines for
handling chemical and biological waste are posted in the lab, and individual waste
containers are clearly labeled to prevent inappropriate mixing. Learn to distinguish
between containers for non-hazardous waste (trash), glass waste (gray buckets at each
TA bench), biological waste (green buckets lined with an autoclave bag, located
throughout the lab), needles and syringes (beige ‘sharps’ containers at each TA bench),
solid chemical waste, including filter paper (cardboard box under the waste bench),
flammable solvents, and acid, base, or oxidizer waste (containers located in the hoods).
If you are not sure about how to dispose of something, ASK.
Check that each chemical waste container is labeled as “Hazardous Waste” and
has a tag that lists its contents.
Replace the cap or cover on the waste container as soon as you have finished
depositing your waste into the container.
Dispose of solid chemical waste and filter paper into the solid waste container.
Dispose of solid biological waste in the autoclave bags in green buckets located
throughout the lab.
Dispose of liquid chemical waste into the properly labeled waste bottle and recap
the bottle tightly as soon as you are done.
Dispose of needles and syringes into the “sharps” biohazard containers under
the TA benches.
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Report full, smoking or foul smelling waste containers immediately.
Material Safety Data Sheets (MSDS) are available in the lab in accordance with Federal
“Right to Know” laws. They provide information about the physical and chemical
properties of chemicals used in the laboratory such as toxicity, flammability, and special
precautions to be aware of in case of spills, exposure, and incompatibilities with other
substances.
The goal of chemical biology research is to collect data from which new insight and
theoretical understanding can be developed. Very little insight is likely to arise from
messy, incomplete, or incoherent records, so you must take care to make your notebook
as detailed and accurate as possible. Not only will you facilitate your own analysis of
your results with a thorough, organized notebook, but you will also be recording
information that might be of use to future investigators.
"The guiding principle for note-keeping is to write with enough detail and clarity that
another scientist could pick up the notebook some time in the future, repeat the work
based on the written descriptions, and make the same observations that were originally
recorded. If this guideline is followed, even the original author will be able to understand
the notes when looking back on them after considerable time has passed." (From Kanare,
H. M. Writing the Laboratory Notebook; American Chemical Society: 1985, p. 1.)
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• Use a blue or black ballpoint pen and write firmly and legibly.
• Use past tense.
• Strike through mistakes with a single line. Your notebook does not need to be a
polished work of art, and you should get in the habit of writing directly in the
notebook as you work.
• Clearly label experiments and procedures within them with descriptive headings.
• When continuing from one page to another, make sure you write “continued on
page __” at the top of the page and “continued from page __” at the bottom of the
next page.
• Sketches or diagrams to illustrate procedures and equipment may be appropriate.
• At the close of each lab period, staple together the carbon copies of your lab
notebook pages, check to make sure that your name is on each page, and submit
them to your TA
At the beginning of each lab period, you are required to hand in an Experimental Plan to
your TA. The purpose of preparing this plan is to help you come to lab prepared to work
efficiently, having thought through your experiments and the necessary preparation for
each step. Always start by reading relevant sections of the lab manual, textbook or
literature references, and the appropriate protocols. Write your Experimental Plan directly
into your notebook. Include as many of the following sections as apply.
b. Objective: Think about what you accomplished during the last lab period and
what you aim to do in the lab period for which you are preparing the plan.
Because of the day-to-day unpredictability of laboratory research (part of its
charm), it is very important that you spend some time after each lab period, and
before the next one, reflecting on what happened and what you learned. Think
about the following questions with respect to your progress in the lab: what did I
learn? What new questions can I ask? What do I need to do to answer them? What
problems arose and how can I attempt to solve them? These reflections should be
succinctly presented in the Objective section of every Experimental Plan.
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intended as a guide as you work, but keep in mind that sometimes research takes
unexpected turns and you will need to revise your plan during the lab. Your TA
will be a valuable consultant when these occasions arise.
e. Teamwork: if you will be sharing equipment with other students, consult with
them before lab begins. Organize your time so that everyone can keep making
progress, and to avoid extensive waiting in line. This kind of efficiency is a
learned habit, one that you will develop over time. If you are not sure what you
might be doing at any time, ask your TA for guidance.
What you write down while you are in lab is the most important part of your lab
notebook. Learn to keep detailed notes as you go. Memory is not sufficient, and jotting
notes on a paper towel or auxiliary sheet of paper to transpose into the notebook later is
not acceptable. Remember that you need to include enough detail so that someone could
repeat your experiment exactly by referring to your notebook. Please observe the
following guidelines for notebook writing as you perform your laboratory research.
• The in-lab section of your notebook should contain details of all the procedures
you perform and all of your observations and data. Try to write with brevity and
legibility. Tables, sketches, and diagrams can be useful.
• You only need to record a technique in excruciating detail the first time you
perform it. After that, make note of its specific application and any modifications
from the procedure you recorded initially.
• Record the actions you take and the observations you make in the order in which
they occur.
• Record calculations in your notebook, clearly showing the formula used and
taking care to include units.
• For data generated on an instrument (the HPLC, the UV-Vis, Mass Spectrometer,
CD, etc.), keep copies of all spectra and other printouts taped in your notebook.
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• Label all spectra with your name, the date, and what is being analyzed. Attach
spectra, chromatograms, photographs of gels, and other data sheets into your
notebook. All figures, spectra, tables, etc. should be given an identification
number by the following convention: your initials, the notebook number, and the
page number. For example, an HPLC trace that Joe P. Student attaches to page 38
of his first notebook would be coded JPS-I-38.
Your notebook will be graded on thoroughness, the quality of your data, clarity of
experimental details, your ability to plan and execute research experiments, and how well
you demonstrate an ability to interpret data and draw conclusions from it. Organization
and legibility will be taken into account.
In general, students will be graded on their understanding of the techniques they are
learning to perform, the quality of their experimental design, laboratory notebooks,
experimental data, and their ability to draw conclusions from results and place them in a
scientific context. There will be four quizzes on material presented in the Friday lectures,
a written proposal for future experiments, and a final report in the format of a short
Journal of the American Chemical Society article for the final exam.
Quizzes are given at the beginning of a Friday lecture period according to the schedule.
They will cover material presented in the previous lecture periods. If you miss a quiz, you
must arrange with the TA in advance to make it up. Make-ups for cases without advance
notice require a Dean’s excuse.
The TA will collect pre-lab assignments at the beginning of every lab period. Late pre-
lab assignments are not accepted and are worth zero.
Laboratory notebook pages (the carbon copies) are turned into your TA at the close of
each lab period and are evaluated as described in Section 8 above. Laboratory
technique will be subjectively assessed by your TA, who will take into account evidence
of preparation, understanding of techniques and concepts, efficiency, safety and waste
management practices, and courtesy toward others.
Each student will develop a proposal for future experiments at the end of the semester.
It should be no longer than five pages in length and must include a summary of the
semester’s results and a detailed description of experiments designed to build upon, or
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further refine, those results. Include sequences, structures, and an experimental plan as
appropriate.
In lieu of a final exam, students will write a final report presenting results from the
semester’s research in the format of a short J. Am. Chem. Soc. article. See Section 10
below for detailed information about how to write this report.
Letter Grades
Letter grades are based on the total percentage of points earned. If necessary, your TA
can scale the grades at the end of the semester. Always keep all of your graded work in
case there is a mistake made in your grade. Letter grades are assigned at the end of the
semester. You can get your grade from the Registrar as soon as it is posted online.
You are required to submit a final report by (date) in the style of a short article from J.
Am. Chem. Soc. All the information you need to help you prepare your article can be
found in this section of the manual, but you may also wish to view the instructions for
authors posted on the JACS website, which is accessible through Yale IP addresses
(https://paragon.acs.org/paragon/application?pageid=content&parentid=authorchecklist&
mid=ag_ja.html&headername=Author%20Information%20-
%20Journal+of+the+American+Chemical+Society).
This journal requires that manuscripts be presented “with the utmost conciseness
consistent with clarity.” Your report should be as brief as possible while allowing
adequate treatment of your results and conclusions. Each paper should contain the
following:
• A descriptive title and list of authors (those who contributed intellectually to the
work).
• A paragraph or two to provide background that will orient the research into a
larger scientific context (what experimental findings informed and inspired your
particular research questions?).
• A description of the question you are asking and the methods you are using to
address it.
• An outline of the experiments performed and the results obtained therein. For
each experiment, clearly state what was being investigated and how the results
provide relevant information.
• To present your results, use figures as appropriate: molecular structures, amino
acid sequences, representative spectra, etc.
• For results that don’t make sense, suggest possible explanations (an excellent
thinking exercise) and ways to test them.
• Summarize your results and present your conclusions. What did you learn?
Provide an opening for future experiments, and suggest a direction for
continuation of the research.
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• Include references as numbered footnotes.
Submit your paper in duplicate along with a CD containing your report, all figures, and
all of your data files from the semester.
Each team of four students will be led by one TA who is an expert in the research project
for that sub-group. Think of your TA as the most valuable resource you have for your
research. He or she will not only supervise your progress, provide experienced technical
assistance, and help you troubleshoot when necessary, but will also serve as a mentor to
your research. Your TA will help you analyze your results and determine what to do next,
thereby directly training you in how to approach and execute research. In addition, your
TA can help you keep up-to-date with the latest developments in your specific research
area by alerting the sub-group to relevant newly published articles.
Sub-group 1: The β-peptide project: What is the effect of salt bridge structure on
14-helix stability?
page 16
Sub-group 3: The phage display project: Can we identify miniature proteins that
bind human MDM2 with high affinity?
page 17
Chemical Biology Laboratory
Project Description
By studying the in vitro and in vivo interactions between rationally designed molecules
and biological macromolecules, we can increase our understanding of the structural and
energetic features of vital cellular events. There is widespread interest in chemistry and
biology in the development of non-natural, functional polymers that mimic, and perhaps
even improve upon, the recognition properties of their natural counterparts. Foldamers is
a term coined by Samuel Gellman (University of Wisconsin at Madison) for “any
oligomer that folds into a conformationally ordered state in solution, the structure of
which is stabilized by a collection of noncovalent interactions between nonadjacent
monomer units” (from Hill et al, see references below). β-peptides represent one class of
foldamers useful for the design of biomimetic structures. These non-natural polymers are
composed of β3-L-amino acids, analogs of natural amino acids that are substituted on the
third carbon. Though they deviate in geometry and side-chain placement from the α-
helix, β-peptide helices have generated interest recently in part because of their surprising
diversity and ability to form compact, stable folds. One such conformation is the 14-
helix, which forms hydrogen bonds between an amido hydrogen and a carbonyl oxygen
to form a 14-atom bonded ring. These molecules form three-sided cylinders (see Figure
1), with side chains lined up along the vertical axes.
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understanding of β-peptide folding will help to further explain molecular assembly and
protein folding, in addition to providing strategies for the design of ligands for
pharmaceutically relevant targets. Recently, coworkers in the Schepartz group have made
progress toward this goal by designing β3-L-amino acid oligomers that form stable left-
handed 14-helices in water. Earlier generations of water-stable β3-peptides required
extensive intramolecular salt bridging and limited the chemical diversity of the side
chains. Our model peptides minimize these requirements by stabilizing the 14-helix
macrodipole. These results raise questions about the relative roles of conformational
entropy and optimal geometry in salt-bridge stabilization of 14-helices.
How does varying the length of the electropositive or electronegative side chains on the
salt-bridging face of the β-peptide affect the stability of the 14-helix? Each of the four
students in this sub-group will independently study a different eleven-residue β3-peptide
based upon a molecule previously characterized by the Schepartz lab. These molecules
will use β3-L-amino acid substitutions to vary the length of side chains along the salt-
bridging face of the 14-helix while maintaining stabilization of the helix macrodipole.
These experiments will provide insight into the geometry of the electrostatic effects
stabilizing these structures, and this knowledge may be applied to optimize β3-peptide
structures toward even greater stability. The β3-peptides will be synthesized using solid
phase methods, purified by HPLC, analyzed by mass spectrometry, and characterized by
circular dichroism spectroscopy and analytical ultracentrifugation. By the end of the
semester, students will be able to analyze the results individually and collectively to
orient their findings within the larger scientific context. Based on these results and the
conclusions they draw from them, students will propose logical future experiments for
the project.
1. HPLC purification
2. Amino acid analysis*
3. Mass Spectrometry of proteins
4. Circular dichroism to determine secondary structure
5. Circular dichroism melting experiments
6. Circular dichroism variations (H20 vs. methanol, pH-based or salt-based screening
of electrostatic interaction, etc.)
7. Sedimentation equilibrium*
8. β3-amino acid synthesis (if time allows)
*out-sourced
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Sub-group 1 Background Reading (link to main reading list)
Journal articles: The majority of these articles are available online to Yale IP addresses.
Papers authored by the Schepartz group and review articles are the best starting place.
Your TA will alert you to especially useful references throughout the semester.
page 20
Deciphering Rules of Helix Stability in Peptides C. A. Rohl, R. L. Baldwin, Methods in
Enzymology 1998, 295, 1-26.
Suggested search terms for this project: β -peptides, foldamers, 14-helix, helix
stabilization, papers authored by S. H. Gellman
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Semester Schedule for Sub-group 1
Required preparation by TA:
4 crude β-peptide syntheses
2 purified β-peptides (as backup; can be material you have on hand)
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Monday (Friday Schedule) QUIZ 4
April 26
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Sub-group 1: Structure-function relationships in 14-helices
Proposed Experimental Flow Chart
Confirm purity of β-
peptide by
reinjection on HPLC
Analyze purified β-
peptide variant by aa
analysis, mass spec
Characterize β-
peptide variant by
sedimentation Characterize β-
equilibrium peptide variant by
circular dichroism
Interpret results;
design a subsequent
β-peptide to
synthesize/analyze
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Chemical Biology Laboratory
Sub-group 2: Analyze the paralog specificity of miniature proteins that bind Bcl-2
and/or Bcl-XL. (return to Contents)
Project Description
Virtually all events in biology are controlled at some level by protein-protein interactions,
and many of these interactions are stabilized by an α -helix at the interface where
recognition occurs. Removing the α-helix from the context of a native protein fold
typically destabilizes the α-helix and destroys its function in folding and recognition. The
Schepartz lab at Yale has pioneered an approach called protein grafting that circumvents
this problem, allowing the design of molecules – miniature proteins – that bind protein
surfaces with high affinity and selectivity and inhibit the formation of protein-protein
interactions. In protein grafting, those residues that comprise the recognition surface on
the α−helix (the functional epitope) are substituted onto the solvent-exposed α-helical
face of the small yet stable protein avian pancreatic polypeptide (aPP). This procedure,
often in combination with molecular evolution, identifies miniature protein ligands with
high affinity and specificity for macromolecular targets. aPP is a 36 amino acid peptide
whose structure contains an α-helix joined by a type I β-turn to a type II polyproline
helix. Because it is small and exceptionally stable, aPP provides a versatile scaffold for
the miniaturization of
proteins employing an α-
helix in macromolecular
recognition.
page 25
protein in the Bcl-2 family, binds to Bcl-2 and Bcl-XL through a 16-residue sequence
known as a Bcl homology domain (BH3 domain). Selected molecules that bind Bcl-2 and
Bcl-XL have the potential to inhibit of their binding to Bak, a tactic proposed to restore
apoptosis in cancer cells. We believe that analysis of the in vitro and in vivo interactions
between miniature proteins and members of the Bcl-2 family will deepen and broaden
our understanding of the structural and energetic components of protein-protein
interactions in general, and serve as lead compounds in a wide variety of bioengineering
and proteomics applications. Previous coworkers in the Schepartz lab have taken the first
step toward this goal by developing miniature protein-binding proteins that are highly
potent and specific ligands for the human proteins Bcl-2 and Bcl-XL.
Can we identify miniature proteins selective for Bcl-2 and Bcl-XL? In this sub-group, we
will explore the specificity of Bak-based miniature proteins selected for binding
specificity to the human proteins Bcl-2 and Bcl-XL. Each of the four students in the sub-
group will independently express, purify, and characterize a different miniature protein
based upon one previously identified in a phage display selection for Bcl-2/Bcl-XL
binding specificity. They will use fluorescence polarization analysis to determine the in
vitro equilibrium dissociation constants of its complexes with Bcl-2 and Bcl-XL. By the
end of the semester, students will be able to analyze the results individually and
collectively to orient their findings within the larger scientific context. Based on these
results and the conclusions they draw from them, students will propose logical future
experiments for the project.
*out-sourced
page 26
Sub-group 2 Background Reading (link to main reading list)
Journal articles: The majority of these articles are available online to Yale IP addresses.
Papers authored by the Schepartz group and review articles are the best starting place.
Your TA will alert you to especially useful references throughout the semester.
A view to a kill: ligands for Bcl-2 family proteins [Review] S. E. Rutledge, J. W. Chin,
A. Schepartz, Current Opinion in Chemical Biology, 2002, 6, 479-485.
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Combinatorial thinking in chemistry and biology J. Ellman, B. Stoddard, J. Wells,
Proc. Natl. Acad. Sci USA, 1997, 94, 2779-2782.
Bak BH3 Peptides Antagonize Bcl-XL Function and Induce Apoptosis through
Cytochrome c-independent Activation of Caspases E. P. Holinger, T. Chittenden, R. J.
Lutz, J. Biol. Chem., 1999, 274, 13298-13304.
Suggested search terms for this project: Bcl-2, Bcl-XL, miniature protein recognition,
BH3 domain, cancer + apoptosis
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Semester Schedule for Sub-group 2
Required preparation by TA:
4 mini protein clones and optimized expression conditions
page 29
Monday (Friday Schedule) QUIZ 4
April 26
page 30
Sub-group 2: Analysis of Bcl-2/Bcl-XL specificity of mini proteins based on Bak
Proposed Experimental Flow Chart
Characterize purified
mini protein by UV,
aa analysis, MS
Interpret results;
design a subsequent Label purified mini
mini protein to protein with (two)
express/analyze fluorophore(s)
Characterize Bcl-2/Bcl-
XL•mini protein Purify fluorescently
interaction by labeled mini protein
fluorescence polarization by HPLC
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Chemical Biology Laboratory
Project Description
Virtually all events in biology are controlled at some level by protein-protein interactions,
and many of these interactions are stabilized by an α -helix at the interface where
recognition occurs. Removing the α-helix from the context of a native protein fold
typically destabilizes the α-helix and destroys its function in folding and recognition. The
Schepartz lab at Yale has pioneered an approach called protein grafting that circumvents
this problem, allowing the design of molecules – miniature proteins – that bind protein
surfaces with high affinity and selectivity and inhibit the formation of protein-protein
interactions. In protein grafting, those residues that comprise the recognition surface on
the α−helix (the functional epitope) are substituted onto the solvent-exposed α-helical
face of the small yet stable protein avian pancreatic polypeptide (aPP). This procedure,
often in combination with molecular evolution, identifies miniature protein ligands with
high affinity and specificity for macromolecular targets. aPP is a 36 amino acid peptide
whose structure contains an α-helix joined by a type I β-turn to a type II polyproline
helix. Because it is small and exceptionally stable, aPP provides a versatile scaffold for
the miniaturization of proteins employing an α-helix in macromolecular recognition.
The goal of this project is to identify and characterize miniature proteins that bind the
human double minute 2 oncoprotein (MDM2) with high affinity and selectivity and
inhibit the interaction of p53 with
MDM2 (see Figure 1).
page 32
for ubiquitin-dependent degradation. We believe that analysis of the in vitro and in vivo
interactions between miniature proteins and MDM2 will deepen and broaden our
understanding of the structural and energetic components of protein-protein interactions
in general, and serve as lead compounds in a wide variety of bioengineering and
proteomics applications. Previous coworkers in the Schepartz lab have taken the first step
toward this goal by identifying a single miniature protein, p53-05, that binds MDM2 with
modest affinity (Kd = 99 nM), only a factor of two better than an unstructured peptide
containing the p53AD sequence (Kd = 261 nM).
Can we use a phage display experiment to select better inhibitors of the p53•MDM2
interaction? Students on this team will generate four variants of p53-05, each displayed
on filamentous phage, and select for those that bind MDM2 with higher affinity and/or
specificity. Four residues within the α−-helix region of p53-05 will be randomized in this
experiment, and those phage that bind GST-MDM2 with maximal affinity and specificity
will be isolated by affinity chromatography. The miniature proteins displayed by this
phage sub-population will be identified by sequencing the phage DNA and then prepared
using solid phase synthesis. These molecules will be analyzed by circular dichroism to
assess their secondary structure (fraction of α-helix) and by fluorescence polarization
experiments to assess their affinity for MDM2. By the end of the semester, students will
be able to analyze the results individually and collectively to orient their findings within
the larger scientific context. Based on these results and the conclusions they draw from
them, students will propose logical future experiments for the project.
*out-sourced
page 33
Sub-group 3 Background Reading (link to main reading list)
Journal articles: Most of these articles are available online to Yale IP addresses. Papers
authored by the Schepartz group and review articles are the best starting place. Your TA
will alert you to especially useful references throughout the semester.
A miniature protein inhibitor of the interaction of p53 with human double minute 2
R. Zutshi, J. A. Kritzer, A. Schepartz, manuscript prepared.
Design of a synthetic MDM2-binding mini protein that activates the p53 response in
vivo A. Bottger, V. Bottger, A. Sparks, W. L. Liu, S. F. Howard, D. P. Lane, Curr Biol,
1997, 7, 860-869.
page 34
Molecular mechanism of the interaction between MDM2 and p53 O. Schon, A.
Friedler, M. Bycroft, S. M. Freund, A. R. Fersht, J Mol Biol, 2002, 323, 491-501.
Suggested search terms for this project: p53, MDM2 (also p53-MDM2 interaction),
phage display, miniature protein recognition
page 35
Semester Schedule for Sub-group 3
Required preparation by TA: Instead of advance prep, this TA is responsible for tasks to
move the project forward during the semester.
Week Laboratory activities Friday Lecture Topic
Friday (Monday Schedule)
Jan. 17 Orientation for MW lab students
page 36
*During the two-week spring break, the TA will analyze sequencing results to identify interesting
molecules for further characterization. He will prepare crude syntheses of two or more of these
selected molecules for purification and analysis by the students upon their return from the break.
page 37
Sub-group 3: Development of mini protein inhibitors of the p53•MDM2 interaction
Proposed Experimental Flow Chart
Identification of
selected sequences;
digests, send for
sequencing
Analyze sequencing
results; synthesize
selected molecules
Purify synthesized
peptide(s) by HPLC;
characterize by UV, aa
analysis, mass spec
page 38
12. Keeping up with the Literature (return to Contents)
Conducting research will be unlike any other laboratory experiments you have performed
so far in several ways. First, the results are unknown and await your discovery. Second,
you are participating in an active research field where many other scientists at Yale and
other institutions play a role. The scientific community values collaboration and strives
for broad dissemination of new findings. Just as you will draw upon the published
experimental results of others to guide and inspire your research, others may also benefit
from learning about your results. Reading articles written by others who are working on
scientific questions related to your own can alert you to new techniques, new approaches,
and maybe even lead you to propose a new approach of your own. Whether or not the
work you do this semester develops into a publishable article, a poster presentation, a
research talk that you can share with others in the department, or all of these things, your
contributions to the field are interesting beyond the scope of the class.
Because research is ongoing and dynamic, there are always discoveries being made and
new things being reported to the scientific public. How can you tap into this constantly
evolving body of knowledge? Since keeping up with the voluminous amount of research
published in the scientific literature is impossible, you’ll want to start to develop an
efficient strategy for keeping up with the more focused area of science that is relevant to
your research interests. Below are some suggested starting points and links for literature
searches. Along with the recommended background reading, each sub-group project
description has a list of key words, which are useful for searches.
Try to set aside some time each week to explore the literature via these gateways. At first,
you might be overwhelmed or unable to easily pinpoint relevant materials. Over time and
with diligent practice, you will become familiar with searching the literature and learn
how to find relevant material in a time-efficient manner.
http://www.library.yale.edu/science/subject/chemistry.html or
http://www.library.yale.edu/science/subject/biology.html
These Yale library links provide a set of discipline-specific resources. A particularly
useful site, accessed via the chemistry page, is the Web of Science, which contains online
journals available to Yale IP addresses. There are also other lists of electronic journals
and links to campus libraries, departments, and reference help.
http://paris.chem.yale.edu/journals.html
This link, part of the Schepartz lab website, provides links to many electronic journals of
particular interest to the chemical biology researcher.
page 39
13. Protocols for Chemical Biology Laboratory Techniques (return to
Contents)
page 40
Affinity Chromatography: Purification of GST Fusion Proteins
by Heather Volkman
adapted for Chemical Biology Laboratory by J. Frederick
(return to Contents)
I. Introduction
Affinity chromatography is one of the most selective types of
chromatography, and it can be a very useful technique for
protein purification. It employs a specific interaction that takes
place between one kind of molecule in the solute and a second
molecule that is immobilized to the stationary phase. The high
affinity binding that occurs between protein molecules and
their specific ligands can be exploited by this technique.
Examples are histidine binding to metal ions, and glutathione-
S-transferase binding to glutathione, as will be further
discussed in this protocol.
Starting with the supernatant of the cell lysis, there are two steps to GST fusion
protein purification. First, the GST fusion protein is separated from all other proteins by
running the supernatant over a glutathione column; the GST fusion protein binds to the
glutathione column and all other proteins are washed away. The GST protein is then
eluted from the column with glutathione. Second, the eluted GST protein is run over a
Nap10 column to remove the glutathione, resulting in a very pure sample containing only
the GST fusion protein.
page 41
Solutions to prepare
Buffer A (for glutathione column)
This buffer is specific to your protein and is usually specified in the literature describing
the fusion protein's purification. Examples include:
page 42
1. Add 1.33 mL 75% glutathione sepharose slurry to column (both 75% slurry and
column are provided in kit).
2. Drain the column of its storage buffer.
3. Wash column 3 to 5 times with 3 mL Buffer A.
4. Add 1 mL of Buffer A to the sepharose. Mix so that beads are suspended and then
add the slurry to the lysis supernatant in 50 mL orange-cap tube.
5. Wash column with an addition 1mL Buffer to remove any remaining sepharose and
add this to the tube.
6. Incubate sample overnight at 4ºC with shaking or rotation.
Evaluation of purification
At this point you will want to run a Phast Gel on the following fractions:
• starting material (lysis supernatant)
• flow through
• wash1
• wash 2
• wash 3
• eluent 1
• eluent 2
• other eluents
page 43
Removal of glutathione on a Nap-10 column
1. Equilibrate column with 3 volumes of storage buffer.
2. Add 1 mL of eluent from glutathione column, collect flow-through and save it for
step 4. This is the void volume and should not have any protein in it.
3. Add 1.5 mL of storage buffer to column and collect flow-through.
4. Do "dot blot" test (blot filter paper with void and protein fractions, then stain with
Coomassie blue) to ensure that your protein is in the 1.5 mL fraction.
5. Use a new Nap-10 column for each 1mL of eluent. When finished, combine fractions
with protein and run a Phast Gel to check for purity.
If the protein is desired without GST attached to it, and there is a cleavage site
built in to the fusion between GST and your protein, you can use a protease to remove the
GST. The following protocol describes cleavage with thrombin using the Novagen
Thrombin Kit.
Thrombin is an endoprotease that cleaves at the sequence Leu-Val-Pro-Arg-↓-
Gly-Ser. There are two ways to accomplish cleavage. The first (and most common)
method involves carrying out cleavage while the GST fusion protein is still bound to the
glutathione column. This method is excellent if you are only interested in recovering your
protein, because after cleavage the GST is still bound to the glutathione and the protein
elutes by itself.
If you need to recover pure GST as well, purify the sample as described above,
then carry out the thrombin reaction to completion in a tube. Run the completed reaction
back through a glutathione column as described above using 1x Thrombin Buffer as
Buffer A. Flow-through will contain your protein plus thrombin, and then you can
remove thrombin as described below. Finally, you can elute GST from column as
described above. Time and amount of thrombin required for cleavage reaction is
dependent on the protein. You may want to optimize the reaction conditions on a small
scale first, starting with a general estimate of 1 unit of thrombin per mg of target protein.
Solutions to prepare
3x Thrombin Cleavage Buffer
60 mM Tris pH 8.5
300 mM NaCl
1 mM CaCl2
page 44
Cleavage on glutathione column
1. Add 1.33 mL of 75% glutathione slurry to column and allow to settle.
2. Drain column.
3. Wash 3x with 5 mL 1x Thrombin Cleavage Buffer.
4. Add sample in 1x Thrombin Cleavage Buffer 2-3 mL at a time. Load and incubate
column in batches if sample volume is bigger than 3 mL.
5. Incubate and rotate/shake for 1 hour at 4ºC.
6. Centrifuge to pack column and save flow-through.
7. Wash column 3x with 1x Thrombin Cleavage Buffer and save washes.
8. Add biotinylated thrombin in 2 mL 1x Thrombin Cleavage Buffer (~1 unit/mg
protein). Incubate and rotate/shake for 2 hours at room temperature or 4ºC, depending
on robustness of target protein.
9. Remove a 20 µL aliquot from the slurry.
10. Spin down aliquot and use supernatant to run Phast Gel to determine extent of
cleavage. At this time you can also run flow-through and washes on the gel.
11. If necessary, incubate overnight and/or add more thrombin
12. When complete cleavage is verified, collect protein by centrifugation. Protein will be
in flow-through.
13. Elute GST as per GST purification protocol (if desired).
It may also be necessary to test cleavage at various temperatures between 4ºC and
37ºC. Once the appropriate conditions are found, scale up the reaction. For more
information on factors affecting cleavage, refer to the instructions provided with the kit.
page 45
Additional information (products, handbooks and instructions as pdf files, etc.)
can be found on the Amersham website at the following address:
http://www1.amershambiosciences.com/aptrix/upp01077.nsf/Content/Products?OpenDoc
ument&parentid=366157&moduleid=38861.
page 46
Agarose Gel Electrophoresis
by Kamil Woronowicz
adapted for Chemical Biology Laboratory by J. Frederick
(return to Contents)
I. Theory
Gel concentration
The concentration of agarose in the gel can be fine-tuned to achieve optimal
separation for a specific range of sizes. The general equation for the relationship between
electrophoretic mobility (µ) and the gel concentration (C) is:
log µ = log µo – Kr C
where µo is the “free” (matrix-free) electrophoretic mobility, and Kr is the retardation
coefficient (a scaling factor) which is related to the properties of the gel and the size and
shape of the migrating molecules (see Maniatis, p. 6.5). A plot of log µ versus C is called
a Ferguson plot and can be used to optimize gel concentration for difficult separations. A
steep slope on such a plot (large Kr) usually indicates a larger molecule, so that sieving
effects become more pronounced at higher gel concentrations. A higher intercept (large
µo) usually indicates a more charged molecule.
For the typical DNA separation experiment, however, this simple chart is
sufficient for selecting a gel concentration:
page 47
0.3 5,000 – 60,000
0.6 1,000 – 20,000
0.7 800 – 10,000
0.9 500 – 7,000
1.2 400 – 6,000
1.5 200 – 3,000
2.0 100 – 2,000
Usually 1 to 2% gels are used for detecting plasmids (several kb long) or their
fragments (ie. from digestions). For resolving much shorter DNAs, use polyacrylamide
gel electrophoresis (PAGE, see separate section). Gels with a lower percentage of agarose
tend to be flimsy, so if you do use them run them at low temperature (4ºC).
Agaroses
There are a few different types of agarose available. For analytical purposes, such
as running digested plasmids to see whether a ligation was successful, you can usually
use agarose from USB. However, if you want to recover your DNA and/or perform some
in-gel reactions, you should use the low melting agaroses (the NuSieve GTG, etc). These
specific agarose protocols are usually provided with the reagent and are available online.
Equipment
To run a gel you will need the following:
1. Two 1L orange cap bottles.
2. 250 mL flask
3. Volumetric cylinders
4. Spatula
5. Gel casting tray
6. Gel combs
7. Tape
8. Electrophoresis tank
9. Power supply and cables
The first six items are used to pour the gel, and the last three are required for running
the gel.
Buffers
There are several buffers that can be used. TAE is typically used, but TBE and
others can be used also (again, see Maniatis). Making a stock of 50x TAE for yourself
saves time and prevents variations in salt concentration from gel to gel. Also, make or get
0.5 M EDTA ahead of time and adjust pH to 8.0 (it can be somewhat time consuming).
50x TAE
242 g Tris-base
57.1 mL Acetic Acid, glacial
page 48
100 mL 0.5 M EDTA
Filter
6x Loading Dye
0.25% Bromophenol blue – BB— (or tiny amount on the spatula tip)
0.25% Xylene cyanol FF –XC— (or same as BB)
15% Ficoll
120 mM EDTA (240 µL of 0.5 M EDTA in 1 mL total 6x loading dye)
Note: Very little loading dye is used; 1 mL of 6x dye should last a long time!!
page 49
3. Add 1 µL loading dye per 5 µL sample (because the dye is 6x).
4. Add samples:
a) Loading 100-500 ng of DNA per lane is usually sufficient.
b) Total sample volume should be from 10-35 µL (depends on the gel thickness
of the gel and well size used).
c) One of the samples should be a marker that contains DNA fragments of
known lengths that are in the range of your samples.
5. Connect the tank to the power supply:
a) Set the voltage at ~150 V. The passage of current will produce bubbles at the
electrodes. Also, flipping the display switch to mA should show you a value
(usually 2 or 3 digits). If you have no current, check the connections. The
samples will migrate towards the “+” electrode.
b) Watch the gel carefully in first couple of minutes to ensure that the dyes are
migrating in the correct direction. If they are not, turn off the power, switch
the electrodes and turn the power back on. The gel should still come out
reasonably well.
6. Run for about an hour or until the faster dye (BB) migrates most of the way through
the gel. You can monitor the progress of the DNA directly (if the EtBr was added) by
shining UV light on the gel as it is running. Just be careful with the UV lamp.
Staining a gel
If you did not add the ethidium bromide earlier, you will need to do so before you
can visualize it. The advantage of staining it after running is reduced probability of DNA
damage and perturbed migration. The disadvantage, however is that you can’t visualize
the DNA directly during the run. If you didn't add EtBr, put the unstained gel in a
container and pour some TAE buffer (you can reuse the one from the tank you just used
to run the gel) just enough to cover the gel, and add ~50µL EtBr (from 10 mg/mL stock
solution). Incubate for about an hour with mild shaking.
If you want to take a picture of your gel and save the image:
page 50
1. Make sure you have a directory on the E: drive on Pompeii (that’s the computer
next to the STORM). Your TA will create one designated for Chemical Biology
students.
2. Place the digital camera (with its black “hood”) onto the gel so that the four
corners of the hood align with the marks on the transilluminator.
3. Open Adobe Photoshop on Pompeii.
4. Go to File Import and click on TWAIN_32. You’ll get a window called “Kodak
DC120 Digital Access (Twain Acquire)." Click “Camera Functions” and set the
following:
• Single Spot (Auto Focus)
• Best (Quality)
• Flash OFF
• Shutter speed to 1/2 second (in the Manual Exposure box); this is a good
starting point as it works for most gels; you can optimize it for your gel, but
keep in mind that you can bring out a lot of details in image processing later.
• Click “Update Camera”. **VERY IMPORTANT** Settings will not take
effect unless you update camera. Update after you have made all desired
changes.
• Close the Camera Functions window.
5. Turn on UV. Click on “Take a picture!” and wait; you should see a small picture
of your gel. If it is all black, you forgot to turn on the UV. If it is faint, you can
change the shutter speed, or you can try to see whether you’ll be able to recover it
in Photoshop (usually you can). If that doesn’t work, double check all previous
steps and try again.
6. Now click on “Transfer picture” and wait for the picture to transfer to Photoshop.
When it does, close the “Kodak DC120” window.
7. Go to Image Mode and click on Grayscale; click OK (discard color
information). You don't have to make it black-and-white, but it will make the file
smaller, and only need to determine the positions of the bands.
8. Go to Image Adjust and click on Auto Levels. This will work great most of
the time. If you still don’t like the way your gel looks, you can change contrast
and brightness manually (under the Image Adjust menu).
9. Also, invert the image so that the bands are black and the gel is white/gray. This
will save the ink when you print.
10. Save the image in your directory on the E: drive.
11. Print the image to include in your lab notebook. Remember to always label each
gel image with the contents of each lane so you know what you are looking at.
page 51
Recombinant Protein Expression and Purification
by Alexis Kays Leonard
adapted for Chemical Biology Laboratory by J. Frederick
(return to Contents)
page 52
Phast gel sample preparation for 1 mL samples from growth: First pellet cells in a
microcentrifuge and carefully decant the supernatant. Then resuspend pellet in 200 µL
2xSDS buffer and heat shock the cells for 2-5 minutes at 95ºC. Finally, run samples on
Homo-20 or Homo-12 phast gel to verify that K97C was not expressed before addition of
IPTG and was expressed after addition of IPTG. You may have to dilute the phast gel
samples to be able to see bands clearly.
1x Ranish Buffer
30 mM Tris-HCl, pH 7.5 at 25ºC
10% glycerol
50 mM KCl
1mM EDTA
2mM DTT (Boehringer Manheim), always added immediately before buffer is used!
page 53
to 2. yTBP elutes around 360 mM KCl (around fractions 17-20). Monitor fractions for
protein concentrations with the UV monitor on the FPLC. Check fractions showing
protein concentration on phast gel (or check all fractions on a Coomassie-stained filter
paper "dot blot"). For the phast gel, mix 3 µL of fraction with an equal amount of 2x SDS
buffer, run a HOMO-20 phast gel. Select TBP-containing fractions.
Concentrate/buffer exchange fractions in Centriprep-10 concentrators at 4ºC.
Reduce KCl concentration to < 100 mM by buffer exchanging with 1x Ranish. Reduce
total volume to less than 5 mL.
Expression optimization
The following is a list of different factors in recombinant protein expression that
can be altered to achieve higher expression of the protein.
1. Transformation
Different cell lines can be used for protein expression. The general cell line used
in our lab is Bl21 (DE3) cells, which can be purchased from Novagen in a competent
form. We also currently have in the lab BL21 (DE3) pARG cells which contain a plasmid
encoding the eukaryotic arginine tRNA that is not naturally present in E. coli cells. There
are many other variations on expression systems as well.
2. Small Growth
A small growth (5-10 mL per 1 L culture) is typically started 24 hours or more
after the transformation of the plasmid into the cells. (Often a transformation is done one
afternoon, and plates are incubated overnight, then the small growth is started the next
afternoon and incubated overnight.)
page 54
I have found greater success in starting the small growth 12 hours after the
transformed cells have been plated. The small growth often takes 3-5 hours to become
cloudy when the colony used to start it is fresher, and the expression of the recombinant
protein is more robust as a result of fresher cells.
3. Large Culture
Try inoculating the large growth (usually 1 L growths in 4 L flasks) when the
small growth first becomes cloudy. As I stated above, keeping the cells multiplying at a
healthy rate seems to result in the best expression.
4. Temperature of Growth
Some proteins may be less stable and therefore will need to be expressed at a
lower temperature. Try expression at 30 ºC rather than 37 ºC. This will greatly increase
the time required for growth, but it may be a more stable environment for the
recombinant eukaryotic protein in a prokaryotic environment.
6. IPTG concentration
With IPTG, sometimes less is more and other times, brute force (lots of IPTG) is
necessary to get acceptable expression of recombinant proteins. Usually, 0.4 to 1 mM
final concentration of IPTG in your large growth is a good range to test.
7. OD600
If aggregation is a problem (i.e., you get expression but see little or nothing when
you purify your protein), perhaps inducing the expression of your protein earlier may
help reduce the aggregation. If you see no expression at all, perhaps your cells need to
grow longer before recombinant protein expression can be induced. A good range to test
is OD600 = 0.6 – 1.0 (log phase growth).
9. Troubleshooting:
1. Do you need to use a protease inhibitor?
2. Are you truly meticulous about keeping cell cultures / cell pellets / protein
solutions on ice when they are not growing? You should be!!!
3. Do you need to increase the volume of the growth to increase expression (500 mL
– 5L)?
4. Are you achieving complete cell lysis?
5. Perhaps a protein cannot survive a PEI or ammonium sulfate precipitation –
perhaps it needs to stay in solution?
page 55
When having trouble, ask yourself:
Do you see overexpression in pre-induction? In post-induction?
Do you see protein in significant concentration in any discarded fractions during the
purification process?
page 56
Centrifuges
by Scott Hart and Kamil Woronowicz
adapted for Chemical Biology Laboratory by J. Frederick
(return to Contents)
The following protocol describes the use of the centrifuges in the Schepartz laboratory.
You can apply this information to guide your use of the centrifuges in the Chemical
Biology Laboratory as well.
Each lab within the Schepartz Lab Complex has a microcentrifuge for use with
1.5 mL or 0.5 mL Eppendorf tubes. These microcentrifuges are also used for the
QIAGEN Miniprep kits, etc. – anything that requires a tabletop centrifuge. Use is
amazingly straightforward: place your tubes in the centrifuge in a balanced arrangement,
close the top, and do one of two things: either set the timer for a long run, or press the
button on the front for a moment to simply 'pulse' the tubes. For work that must be done
at 4ºC, there is a microcentrifuge in the deli case in the hot room.
The deli case in the hot room and the deli case near Kamil's bench (room
KCL100) also contain centrifuges appropriate for 50 mL conical tubes. Make sure your
tubes are balanced, cap included, before spinning.
A swinging rotor centrifuge is in Joshua's hood (K102). The rotor for this
centrifuge is appropriate for 15 mL Falcon tubes.
Keep in mind that Speed-Vacs are not typical centrifuges; they are specifically for
drying small samples, and procedures for speed-vacs are outlined elsewhere in this
manual.
For proper use of any centrifuge, keep a few simple things in mind: balance your
sample tubes, clean up any messes you happen to make, and inform your TA of any
problems with the centrifuges.
This is a general overview of the preparative centrifuges available to our lab in the
Kline Chemistry Building. Remember, none of these centrifuges belong to our group, so
be conscientious when using them. Betty Freeborn in the Moore Lab is the person in
charge of them. Always clean up after yourself, and log usage in the appropriate logbook.
Always ask for help from someone who has used the centrifuge you need before starting
for the first time.
For information on appropriate rotors and conversion information from RPG to
RPM, see the Schepartz lab website directory of 'Cool Science Links'
(http://www.paris.chem.yale.edu/links.html), where you will find links to Sorvall and
Beckman rotor caclulators.
page 57
This centrifuge belongs to the Crothers group and is not an untracentrifuge. It is
useful for spinning bacterial broths and working up the cells. The common rotor is the
SGA rotor that accepts bottles with volumes of 250 mL each. This one is ideal if you
have up to three 500 mL broths because a single spin will be sufficient to pellet the cells.
There is also a GS-3 rotor that takes 6 500 ml tubes. It is great for large volumes (as is the
Beckman J2-21 discussed below). When using this centrifuge, remember that there is no
vacuum in the chamber, so you cannot perform very high-speed spins. For the SGA rotor,
and for the GS-3 rotor, you will need to get above 6000 rpm to do your work. To display
the speed in "g" you can flip a switch from rpm to rcf.
Procedure:
1. Place the rotor you wish to use in the centrifuge. Set the temperature and wait for at
least 30 minutes to allow the rotor to cool before you do your run.
2. Pour your sample into at least two bottles that have screw top lids. The O-ring lids
work best to prevent leakage during the spin.
3. Balance the samples to within 0.1 g (including the lid).
4. Place the bottles in the cooled rotor.
5. Attach the lid with the two attachment screws in the direction shown.
6. Secure the top on the chamber.
7. Set the speed.
8. Set the time and begin your run.
9. Remove your samples when the run is complete.
10. Check carefully that the bottles have not ruptured or leaked into the rotor.
11. Remove the rotor from the centrifuge. Turn off the power when finished and leave the
lid open.
12. Clean and dry the rotor thoroughly when finished. Do not use harsh chemicals to
clean the rotors or damage will result. Use soap and water and a teflon brush (in the
Moore's lab across the hall; ask Betty if you can't find it). Place rotor upside down on
the paper towels when finished.
13) Clean all the condensed water that formed in the open centrifuge.
Beckman J2-21
This centrifuge belongs to the Moore Lab and is useful for the same types of runs
as the Sorvall. It will accept several different rotors, including one with a maximum
bottle size of 500 mL, so it is useful for large bacterial growths.
Procedure:
The procedure is the same as for the Sorvall. There is also a vacuum that must
register in the green region of the gauge prior to starting your run. For the rotor that
accepts the 500 ml tubes, use a special crowbar to insert a rotor into the centrifuge; the
rotor is quite heavy and the inside of the centrifuge is narrow. You will find the crowbar
next to the rotor (usually). The crowbar screws into the center of the rotor with the arm
that has threads.
page 58
This is an ultracentrifuge useful for cesium chloride gradients of plasmid
preparations or ammonium sulfate fractionations in protein preparations.
Procedure:
1) Choose your rotor. Only use rotors designed for use in this centrifuge!
2) Cool the rotor by storing at the desired temperature for at least 1 hour.
3) Once the sample/rotor is in the centrifuge, turn on the vacuum and wait until the
chamber is <100 microns. This can take awhile so you do not need to monitor it
constantly.
4) Set the desired speed, time, and brake. Do not exceed the speed ratings of the
rotor!
5) Start the run.
6) When run is complete, follow the above instructions for cleaning and storage of
the rotor.
7) Turn off the instrument.
page 59
Circular Dichroism
by Neal Zondlo (1994), edited by Scott Hart (2001)
adapted for Chemical Biology Laboratory by J. Frederick (2003)
(return to Contents)
Background
The most important physical/optical concept in CD is the idea of circularly
polarized light. In CD, the polarized beam of light can be considered to be composed of
right- and left-handed circularly polarized components. The CD instrument itself uses a
double monochromator to take a beam of light (UV range) and eliminate stray light. The
two monochromators are oriented in different axial directions, which serves to produce
linearly polarized light. That's the simple part. The fancy part is called the CD Modulator.
The linearly polarized light is passed through a quartz crystal that has been subjected to
mechanical stress, producing circular polarization within the crystal. Polarized light that
has passed through this crystal is thus modulated to circular polarization.
Now imagine this light passing through an optically active substance. When the
light passes through an optically active material, its two components (left- and right-
handed circularly polarized light) are absorbed to different degrees. This difference in
absorbance of the two forms of light is called circular dichroism.
The light that has passed through the optically active substance shows a net effect
of being elliptically polarized. Much math is involved in truly understanding this, but if
you consider the result of equal portions of left- and right-circularly polarized light as
resulting in a circle (no circular dichroism), consider differential amounts of these types
of light as producing an ellipse. At the cartoon level, this hopefully makes sense. The
molecular ellipticity resulting from this phenomenon is represented by the symbol Q
(theta).
page 60
Figure 1. Representation of circular dichroism spectra for various secondary structures.
Top curve is helix, middle curve is sheet, bottom curve is coil. Taken from Greenfield
and Fasman. For more information of diagnostic spectra for various secondary structures,
see: Greenfield & Fasman, Biochemistry, 8(10), 4108-4116, 1969.
Start-up procedures
Spectrometers often require up to an hour to warm up and get to the proper temperature.
Make sure that this takes place before you prepare your samples and materials to do the
page 61
experiment so you can avoid wasting time. Your TA will sometimes take care of this in
advance, but it is your responsibility to think about it.
Sample handling
The specific details of sample handling will vary depending on the exact instrument used.
In general, you first need to decide which size cell you want to use, 0.1 or 1.0 cm. This
decision is influenced by the concentration of your sample (which affects the absorbance:
you need a strong signal, but not too strong to overwhelm the instrument) and the
strength of the CD signal. If either of these gets too high, you will see the dynode voltage
(dynV) rise, and data acquisition may stop. When handling CD cells, you should wear
gloves and use only lens paper to dry or wrap the cells. Use only plastic tips to remove
solutions from the cell. For 0.1 and 1.0 cm cells you will use 150-200 µL and 300 µL of
sample, respectively. Be sure to mark down the size of the cell you used!
During a run you should close the cells with parafilm or the teflon stopper to
guard against evaporation. After the run, remove the sample from the cell, wash
repeatedly with 1% SDS, water and ethanol solutions, and finally rinse with ethanol and
dry using N2, wiping off excess ethanol with lens paper. A useful diagnostic experiment
to try is to run a spectrum, remove the sample and wash the cell, and then return the
sample to the cell to assure that the spectrum remains the same. At the end of the
experiment, clean the cell thoroughly, wrap it well in lens paper, and immediately return
it to the CD supplies drawer.
Depending on your experiment, you will be able to adjust parameters such as wavelength,
temperature, step size, and number of scans. Your TA will help you determine the
appropriate settings for your experiment. Examples of parameters that have worked well
for b-peptide experiments are as follows:
Temperature 25°C
Path length 2 mm
Averaging time 2 seconds
Bandwidth 2 nm
Peptide Concentration80, 40, 20, and 10 mM
page 62
Collect the relevant data and obtain printouts of the spectra to include in your lab
notebook. Be sure to save your data!
Shut-down procedures
Just as there are steps to go through when you are starting up the CD, there are also shut-
down steps to put the CD into a safe stand-by configuration. You will also need to turn
off the power, allow the lamp to cool, and turn off the gas and water supplies. Follow the
specific instructions for the CD spectrometer you are using.
page 63
Cloning, Version 2.0
by Stacey E. Rutledge
adapted for Chemical Biology Laboratory by J. Frederick
(return to Contents)
1. General Considerations
Cloning is mysterious. What works well for one person will, for no apparent
reason, not work well for another person. You have to find things that work well for you,
and stick with them. That said, this protocol is meant to provide you with some general
guidelines.
When you prepare your insert, you inevitably lose quite a bit of DNA at each step.
Therefore, do everything on a fairly large scale (especially when working with libraries).
It may take an extra half hour to set up ten extra PCR reactions or even an extra day to
gel purify more oligos, but if you play it safe, you won’t ever get to the end of your insert
preparation and realize you do not have enough DNA to do ligations, thus necessitating
weeks of more work to prepare more insert.
page 64
It is useful to design your insert so that ligation of your insert into your vector
creates a restriction site which can be used as a positive screen for the presence of insert.
If you are lucky, the sequence you are inserting will contain a single restriction site which
is not contained in your vector. If not, you can use silent mutagenesis to create a unique
restriction site. I have found the program Webcutter (available from our links page) to be
useful in this regard.
Because most DNA inserts are very small (50-200 bp), it is often difficult to
distinguish fully cut insert from uncut or singly cut insert. For this reason, it is useful to
design your predigested insert such that restriction digest on either end will cut off 15-20
base pairs or more.
Oligos can be ordered on an 0.2 or 1.0 µmole scale. They can be ordered from the
Keck center at Yale Medical School (which has always worked well for me) or from
Operon (see Kevin), or other commercial vendors. If you are ordering long oligos for
library construction, Lori (the resident oligo synthesis expert) has the particulars of how
they should be ordered to ensure efficient synthesis.
Purification
Time estimate: One day per gel + overnight for elution + a few hours for drying.
Primers do not need to be gel purified, and can be used directly after desalting.
Longer oligos can be ordered purified (Beware! They are not always very pure!) or they
can be easily purified by denaturing PAGE followed by desalting. I generally resuspend
oligos on a 1.0 µmole scale in 500 µL dH20, add 500 µL formamide loading buffer (For 1
mL: 980 µL deionized formamide, 20 µL 0.5 M EDTA, spatula tip of xylene cyanol and
bromophenol blue). The oligos are heated to >95 °C for 10 minutes, then quick cooled on
dry ice before being loading on an appropriate percentage denaturing acrylamide gel (see
table).
I load 200-300 µL of each oligo in each well (3 wells/comb). The oligos are
excised from the gel (take care to avoid n–1 contaminants!), crushed through a 5 mL
syringe into a 15 mL orange-cap tube, and eluted in 3 volumes TE (10 mM Tris, pH 8.0;
1 mM EDTA) overnight with shaking. The acrylamide is pelleted by centrifugation and
the supernatant carefully transferred to eppendorf tubes. The oligos can be dried in the
speed-vac and resuspended in 1 mL dH2O for desalting.
page 65
Recommended Polyacrylamide Gel Percentages
for Resolution of DNA
Gel percentage DNA size range
3.5% 100-1000 bp
5% 75-500 bp
8% 50-400 bp
12% 35-250 bp
15% 20-150 bp
20% 5-100 bp
Desalting
Time estimate: 1 hour + drying time
NAP10 columns (Pharmacia) are used for desalting. The columns are equilibrated
with three column volumes of water or TE. The load volume for the columns is 1 mL.
Oligos can be eluted with 1.5 mL dH20. The concentration of desalted oligos can be
determined by measuring UV absorbance (A260) and converting this to concentration via
the Schepartz Lab biopolymer calculator. The oligos are dried in the speed-vac and
resuspended to a concentration of 50 µM.
Annealing
Time estimate: 1.5 hours + 1.5 hours for agarose gel
To anneal, equimolar amounts of each oligo (either two long oligos, or a long oligo
and a primer) are mixed, heated to >95 °C for 10 minutes, then slowly cooled to
room temperature. To monitor the success of the annealing reaction, run a 3%
agarose gel with your single-stranded DNA #1 in one lane, DNA #2 in another lane,
and then the annealed DNA in a third lane. Run a DNA ladder (100 bp (NEB) or
φX174 HinfI (Promega)) in another lane for comparison. Note that the DNA ladders
are double-stranded, so your single-stranded oligos will not have the same mobility
as markers of the same length.
Time estimate: 3 hours + 1.5 hours for agarose gel + 0.5–2 hours for optional
cleanup
The primer for primer extension reactions should be 20-30 bp in sequence,
complementary to either the 3´ or 5´ end of the template oligo, and GC rich. Both ends of
the primer should be a G or C base (preferably two in a row). Primer extension reactions
are performed as follows:
Step 1. Annealing
page 66
400 pmol long DNA
400 pmol primer
40 µL 5x sequenase buffer (USB)
200 µL total volume
The reaction should be heated to >95 °C for 10 minutes, and then slow cooled to room
temperature by removing the hot block from the heating apparatus.
Step 2. Extension
To each annealing reaction, add:
2 µL 25 mM dNTP's
2 µL 10 µg/µL BSA
2 µL 100 mM DTT
4 µL 13U/µL sequenase (USB)
The reaction is incubated at 37 °C for 30 minutes, and then incubated at 65 °C for 1 hour
to heat inactivate the sequenase.
One test reaction should be attempted first, and the success of the primer
extension reaction determined by running 5 µL of the reaction on a 3% agarose gel
(single-stranded DNA should be run also, as a comparison.) If the reaction is successful,
multiple primer extensions can be performed.
After primer extension, digests can be performed immediately on the primer
extension mixture (this is the way I have always done it). However, the high salt and/or
high protein concentration in the primer extension reaction may interfere with restriction
digests. If you find this is a problem, you can clean up your DNA in a number of ways:
EtOH precipitation – add 2.5 vol ice-cold EtOH, 1/10 vol 3 M NaOAc, incubate on dry
ice for 30 minutes, spin for 15 minutes, remove supernatant, wash pellet carefully with 1
vol 70% EtOH (room temperature), remove supernatant, dry pellet, resuspend in
appropriate volume of dH2O or TE.
Phenol/Chloroform extraction – add 1 vol 25:24:1 Tris-buffered
phenol:chloroform:isoamyl alcohol, vortex, centrifuge for 10 minutes, transfer aqueous
layer (top, contains DNA!) to different eppendorf tube.
Nucleotide removal kit (Qiagen) or PCR Purification Kit (Qiagen).
Preparing an Insert from Multiple Synthetic Oligos using Mutually Primed Synthesis
(MPS)
Time estimate: 3 hours + 1.5 hours for agarose gel + 0.5–2 hours for cleanup
(optional)
MPS is just glorified primer extension! The oligos to be used in the MPS reaction
should contain a 20-25 bp overlap, should be fairly GC-rich in the overlap region and
should contain G or C bases at either end of the overlap region.
page 67
The oligos can be annealed and extended under the same conditions as those
described above for primer extension (substitute 400 pmol long oligo #2 for primer). The
success of the MPS reaction should be monitored by running 5 µL of the MPS reaction
and each ssDNA on a 3% agarose gel. Multiple reactions should then be performed. If the
MPS reaction is not clean, reaction conditions can be varied. Some things to try are
changing the MgSO4 concentration, eliminating BSA, changing the annealing
temperature, or changing the extension time.
As with primer extension, these reactions can be cleaned up by one of the
methods discussed previously, or can be used directly.
Time estimate: 3 hours + 1.5 hours for agarose gel + 0.5–2 hours for cleanup
To PCR an insert out of a different vector, you need to order two primers (0.2
µmole scale), one for each end. These PCR primers must contain a 20-25 bp region
identical to the 5´ region of the gene to be amplified (5´ primer) or a 20-25 bp region
antiparallel to the 3´ region of the gene to be amplified (3´ primer). The hybridization
portions of the primer should be GC rich and should contain GC clamps at the ends. In
addition, you may want your primers to contain an overhang region coding for restriction
sites, an initiation codon (5´ primer) and/or a stop codon (3´ primer).
The success of the PCR reaction can be affected by a number of things (especially
primer concentration and Mg2SO4 concentration), and thus a number of PCR conditions
should be screened. Note that you need only a very small amount of template DNA (1 µL
of a 1:100 dilution of miniprepped plasmid is sufficient). A good place to start in testing
PCR conditions:
Volume (µL)
Primer 1 (10 µM) 4 4 6 6 8 8
Primer 2 (10 µM) 4 4 6 6 8 8
100 µM Mg2SO4 2 4 2 4 2 4
25 mM dNTP’s 1 1 1 1 1 1
Template DNA 1 1 1 1 1 1
Thermo Pol Buffer 5 5 5 5 5 5
Vent (exo-) Polymerase 2 2 2 2 2 2
50 µl total reaction volume
page 68
5 to 10 µL of each PCR reaction should be run on an agarose gel to determine the
success of the reactions. If the reactions look clean (only your desired product is visible)
the reactions can be cleaned up as discussed above. I find the PCR purification kit
(Qiagen) works very well for this (use 1 column/ PCR reaction). If the PCR does not look
very good, some other things to try are varying the concentrations of primer or Mg2SO4
further, adding BSA, using a different polymerase or changing the annealing temperature.
PCR can also be used to amplify small amounts of any double-stranded DNA
made by any of the previous methods. To do this, you need primers identical to the 5´
region of the insert to be amplified (5´ primer) and antiparallel to the 3´ region of the
insert to be amplified (3´ primer).
Time estimate: As long as it takes! (1–3 days, generally) including gels to monitor
reactions + cleanup of digested oligos (2 hours – 2 days)
Digestion
The NEB catalog should be consulted for buffer requirements and enzyme
compatibility. Some enzyme combinations are compatible, others are not. Note that many
restriction enzymes cut poorly close to the ends of oligos. Thus, for library inserts, it is
essential to prepare A LOT of double-stranded insert, because you will most likely need
to purify doubly cut insert away from uncut or singly cut insert.
If the required enzymes are compatible (BglII/NotI, for example), the DNA can
be cut with both enzymes at the same time. (For library inserts, I generally do
MULTIPLE (5-7) digests in a volume of 50 µL, cutting 10 µg of DNA in each reaction,
with an appropriate volume of enzyme). In general, you want to keep the volume of the
reaction as small as possible while still cutting as close to completion, so that you can
load more DNA across a smaller number of lanes when you purify it, whether on
acrylamide or agarose.
If the enzymes are not compatible (SfiI/NotI, for example), the DNA should be
cut sequentially. I do this by cutting first with one enzyme (again, 10 mg DNA in 50 µL
reaction), EtOH precipitating the DNA after the first digest is complete, resuspending the
DNA in an appropriate volume dH2O (generally 30 µL), and digesting with the second
enzyme (again in 50 µL reactions). Alternatively, you could cut with the enzyme
requiring less salt first, then add salt to the required concentration and cut with the second
enzyme.
Digestion should be monitored along the way. If a digest has not progressed
sufficiently, it is a good idea to add more enzyme to the reaction and continue the
incubation. For sequential digests, I always run an agarose gel to check that the first
digest has gone to completion BEFORE I go on to EtOH precipitate the DNA and cut
with the second enzyme.
page 69
In the end, if the digest appears to have gone to completion, it may not be
necessary to gel purify. In this case, the digested DNA could be cleaned up using the
PCR purification kit, the nucleotide removal kit, or just by EtOH precipitation, or
phenol/chloroform extraction. If this is not the case, gel purification is necessary to purify
your digested insert.
Quantification
The DNA should be quantified by UV prior to ligation. The final concentration of
the DNA insert should be around 10 ng/µL for it to be useful in ligations. For cloning of
libraries, you need at least 1 mg of insert, but more is better (I usually aim for 3 µg total).
For single sequences, 300 ng is probably enough (I usually aim for 1 µg total).
page 70
Time estimate: Overnight + 4-5 hours for maxiprep of vector, 1-2 days for digests
and cleanup
Ligation
page 71
5 µL 2.5 mg/mL BSA
5 µL 100 mM ATP
25 µL 1 M Tris, pH 8
10 µL 0.5 M DTT
50 µL total volume
Electroporation
Time estimate: 2–2.5 hours including plating + overnight for plates
page 72
place of DNA for a negative control, and another plasmid (such as pUC) can be
transformed as a positive control.
Cells are recovered by immediately adding 960 µL of any rich media (+ glucose
for phagemids) lacking antibiotics to the cuvette. The cells can be transferred to 5 mL
falcon tubes and are incubated for 1 hour with shaking at 37 °C. Plate 50-100 µL of the
recovered cells (neat) and multiple dilutions made in media without antibiotics (10-1 to
10-5) on appropriate agar plates (2xYT-AG for phagemid vectors, LB Amp for most other
plasmids) to ensure that that you will be able to pick single colonies and/or determine the
number of transformants. Plates are incubated at 37 °C for 12-16 hours (not longer!). The
remaining cells are mixed with 500 µL of 50% glycerol and frozen on dry ice.
Heat Shock
Time estimate: 1.5-2 hours including plating + overnight for plates
Heat shock competent cells, including BL21(DE3) and many other strains, are
first thawed on ice. 1 µL of DNA is added to 20 µL cells and mixed by gentle stirring
with a pipette tip. The cells and DNA are incubated for 5 minutes on ice, and then heated
to 42 °C for 30 seconds exactly. The cells are incubated for 2 additional minutes
(exactly) and 80 µL of any rich media (without antibiotics) is added to the tube. The cells
are recovered at 37 °C for one hour, with shaking. Plating and controls are performed as
described above for electroporation.
Screen around 10 colonies for single sequences, or at least 20 for libraries, off of
the plate with the highest ligation efficiency. Grow 5 mL overnight cultures in
appropriate media, each inoculated with a single colony, for 15-16 hours. Minipreps of 2
mL of each overnight culture should be sufficient for screening and sequencing. Two
miniprep kits popular in the lab are the Promega Wizard kit and the Qiagen spin kit. Both
are easy to use, and the manufacturer's directions can be followed exactly. Two digests of
each clone should be performed: one with an enzyme whose site is found in the new
insert but not in the vector (this enzyme should cut your clones!), and another with an
enzyme whose site is found in the region of the vector cut out, but not in the new insert
(this enzyme should not cut your clones!). The digests are performed in a 10 µL volume,
with 1 µL enzyme and 5 µL miniprepped DNA for one hour, and then loaded on a 1%
agarose gel. Clones that show the correct pattern of digestion can be sent to Keck for
sequencing. For some people, sequencing is more successful when they follow the Keck
guidelines for sample preparation exactly. Other people find that, in their hands, other
amounts of DNA are more likely to be sequenced cleanly. I use 15 µL miniprepped
DNA, 7 µL dH2O, 2 µL µM primer.
Cloned Libraries
The number of transformants needs to exceed the theoretical diversity of your
library. To calculate number of transformants, determine the best plate for counting
colonies and multiply:
page 73
(# of colonies) × (dilution) × (1000/how much you plated).
For example, if you have 46 colonies on the 10-4 dilution plate (and you plated 100 µl),
you have 4.6 x 106 transformants / 1 µl of ligation. If your diversity is 3.2 x 107 (5
residues randomized), you will have to transform multiple aliquots of your ligations and
possibly do multiple ligations. To guarantee 90% completion of your library, you need a
number of transformants that is at least 2.3 times the theoretical diversity.
The recovered cells from the required number of transformations are combined and
grown overnight in a large volume of 2XYT-AG (the volume required will depend on
how many transformations you need to combine). You want to have a healthy culture
of cells in the morning. From this culture, you can maxiprep your library. In addition,
you should make multiple (~30) glycerol stocks (1 mL cells + 500 µL 50% glycerol;
freeze on dry ice). These aliquots should each contain multiple copies of every
transformant and can be used directly in phage display experiments.
page 74
DNA Technical Information
Adapted by Lori Yang from www.biosyn.com/t_dna.htm
Adapted for Chemical Biology Laboratory by J. Frederick
(return to Contents)
Calculations
Molar conversions
1 µg of 1,000 bp DNA = 1.52 pmol (3.03 pmoles of ends)
1 µg of pBR322 DNA = 0.36 pmol DNA
1pmol of 1,000 bp DNA = 0.66 µg
1pmol of pBR322 DNA = 2.78 µg
Other Information
Resuspension Buffers
1. Sterile Water (dd H2O)
2. TE Buffer (10 mM Tris-HCI, 1mM EDTA) pH 7.5
page 75
Dissolved (-20˚C) = 1 month to 6 months
Dissolved (25˚C) = 1 week to 3 months
DNA Conformations
References
1. Suggs, S., et. al. Proc. Natl. Acad. Sci 78:6613('81)
2. George H. Keller, Mark M. Manak, DNA probes; p 15; M Stockton Press, '89.
page 76
Fluorescence Techniques
by Tanya Schneider
adapted for Chemical Biology Laboratory by J. Frederick
(return to Contents)
I. Fluorescence Quenching
page 77
FRET can also be used to measure protein•protein and protein•DNA binding. In
contrast to a quenching experiment, FRET requires a pair of fluorophores. Here, a donor
fluorophore is excited and, when in close proximity with an appropriate acceptor
fluorophore, transfers energy to the acceptor fluorophore. In this case, increased
fluorescence is detected for the acceptor fluorophore and decreased fluorescence is
detected for the donor fluorophore. One is able to detect binding by measuring the energy
transfer that occurs when the molecules are proximal (usually donor quenching is
quantified for Kd measurements). A successful donor-acceptor pair must have overlap
between the emission wavelength of donor and the absorbance wavelength of the
acceptor. The pair must also be able to transfer energy over the distance that you estimate
to be relevant for your system. The characteristic transfer distance (R0) is known for
common donor-acceptor pairs. Fluorescein and rhodamine are commonly used as a
donor-acceptor pair due in part to the strong signal of fluorescein.
Unlike fluorescence quenching as described above, FRET gives an indication that
the quenched signal of the donor fluorophore is related to an interaction with the acceptor
molecule based on the acceptor signal. However, there are limitations here as well. If the
donor and acceptor fluorophores are not positioned correctly, no transfer will be seen, so
some thought and molecular modeling may be necessary prior to covalent modification.
Also, it is important to make sure that donor quenching is not due to any factors except
for the presence of the acceptor. One can perform control wavelength scans without the
acceptor fluorophore to check this.
Useful references:
1. Molecular Probes catalog or website (http://www.probes.com/).
2. Fluorescence resonance energy transfer. P. R. Selvin. Methods in Enzymology (1995)
246: 300-334.
3. Kinetic studies of Fos-Jun-DNA complex formation: DNA binding prior to
dimerization. J.J. Kohler & A. Schepartz. Biochemistry, (2001) 40, 130-142.
4. Hepatitis B Virus protein pX enhances monomer assembly pathway of bZIP-DNA
complexes. T. L. Schneider & A. Schepartz. Biochemistry, (2001) 40, 2835-2843.
5. Kinetic preference for oriented DNA binding by the yeast TATA-binding protein
TBP. Y. Liu & A. Schepartz. Biochemistry, (2001) 40, 6257-6266.
page 78
Iperp
P= Ipara – + Iperp
Ipara
Iperp
Anisotropy = Ipara – + 2Iperp
Ipara
2P
Anisotropy =
3–P
1. Used for dimerization curve fit in several Schepartz papers, adopted from CD
Kd fit in:
Zitzwitz, J. A, Bilsel, O., Luo, J., Jones, B. E., and Matthews, C. R. Biochemistry,
(1995) 34: 12812-12819.
where Fapp = apparent fraction of unfolded protein at any concentration and Atot is
the total protein concentration, expressed in terms of the monomer.
2. Used for binding curves in several papers, often for polarization data, taken
from:
Heyduk, T. and Lee, J. C. PNAS, (1990) 87: 1744-1748.
1 (K a * AT + Ka * BT + 1)2 − 4K2a * AT * BT
page 79 P = 0.5(AT + BT + K ) ± 2Ka
a
where P = polarization, AT and BT are total protein/DNA concentration, and Ka is
equilibrium association constant. Typically, either A or B would be fluorescently labeled,
and the concentration of the labeled molecule would be kept constant while the other
species varied over a range of concentrations.
The Molecular Probes website contains useful information about fluorophores, including
a helpful (though general) protocol providing guidelines for labeling peptides with thiol-
reactive probes (see http://www.probes.com/media/pis/mp00003.pdf).
page 80
1. Use a lower concentration of both peptide and rhodamine in the reaction (closer to 25
mM) to avoid precipitating modified peptide, rhodamine, or both.
2. Do not use a Nap-10 column - material will get stuck in the column.
3. Use higher concentrations of DMF or other organic solvent (acetonitrile may also
help) to keep rhodamine in solution if necessary.
4. If everything crashes out of solution, try separating precipitate from supernatant – the
precipitate likely contains modified peptide which you may be able to re-dissolve in
DMF and purify on HPLC.
All other fluorophores have proven more challenging in my experience than fluorescein,
so, if possible, use that.
Buffer: 1X PBS may be fine, or your peptide may demand detergents or other
components to stay in solution. Check to make sure that additions to your buffer do not
greatly change the polarization of your labeled molecule alone if you’re doing
polarization – glycerol has a large effect which masks many polarization changes due to
binding.
Equilibration time and temperature: You can check to see when binding has
reached equilibrium by watching polarization or FRET or quenching with time in the
fluorimeter. This will give you an idea of how long you need to wait before taking
measurements. The fluorimeter currently does not have temperature control. Some people
have done 4 ºC measurements by incubating their binding reaction on ice prior to
measurement, then adding the sample to the cuvette and measuring immediately.
page 81
VI. Using the PTI Fluorimeter
The following is a guide to aid you in use of the instrument. This does not replace
personal instruction from a lab member well acquainted with the instrument.
Start-up
5. Sign in and record start time in order to keep track of lamp hours.
6. Make sure that all other components (computer, motor) are turned off, and turn
on the lamp power. The lamp will ignite automatically after a few seconds. Allow the
lamp to warm up for at least 15 minutes at ~60 watts. Set lamp to 70-75 watts after
warm up. **Igniting the lamp with the computer on can cause damage to the
computer. Also, it is better to leave the lamp on if you’re only leaving the system for
an hour or so - ignition is what really wears on the lamp.**
7. Turn on Motor Drive Box, which powers most of the system. Check to see that the
PMT digital readout is set at 1000V (max = 1100V).
12. Turn on computer. Operating software for the PTI system is Felix. In the Felix
program, first choose Configure. Under Hardware, choose Initialize to set the
monochromators. Check to see that the monochromators are actually set to the
values that the computer gives after initialization.
Taking measurements
Wavelength scans: useful for FRET or quenching experiments. Choose Emission
scan under Acquire to bring up a relevant window. Input the desired excitation
wavelength for your sample and the emission wavelengths you wish to scan. I find that
the default settings for step size (1 nm) and integration time (1 sec) are generally
reasonable, but can be changed as needed. Adjust slitwidth as necessary - each turn of the
screw = 2 nm. I find that 8 - 10 nm is fine for fluorescein-labeled samples. Data can be
saved as .txt files and imported onto a Mac using Excel.
Ihv
G factor =
Ihh
page 82
To measure the G factor, set both polarizers to the horizontal position (90º) (Ihh).
Start a time based scan which is the same as your experimental time will be. I generally
collect 1 point/second for 30 seconds. Then, switch the emission polarizer to the vertical
position (0º) and repeat the measurement (Ihv). The G factor is simply the ratio of the two
measurements as described above. To set the G factor, select the G factor curve where it
is listed on the left of the screen and also highlight the scan on the screen. Choose
Polarization under Configure, and click “capture” to set the G factor. For this
instrument, it is usually roughly 0.7. If you are using a low concentration of labeled
sample, it may also be advisable to subtract out background from your buffer by doing
the same measurements with just the buffer in the cuvette. Subtract these hv and hh
measurements from your sample hv and hh measurements before calculating the G factor.
Polarization measurements, as described above, are based on measuring the
sample with the polarizers in two different positions; the excitation polarizer is always in
the vertical position, but measurements are taken with the emission polarizer in the
vertical and horizontal positions. Thus, you will collect two sets of data for each sample,
and then use these values to solve for polarization. Felix will do this calculation from
your two sets of data - simply choose Polarization under Transform and select the
correct data sets as listed in the left hand column. Choosing Average under Math and
highlighting the polarization curve allows one to measure the average polarization over
the time period of the experiment. As with most experiments, at least three independent
sets of data are needed.
Shut down
1. Close Felix and shut down computer after saving data to disk if needed.
2. Turn off Motor Drive Box.
3. Turn lamp down below 60 watts, and then turn off.
4. Sign out and log total lamp hours.
Final notes
The cuvette should be stored with distilled water in it after cleaning. Occasional
careful cleaning with nitric or hydrochloric acid can be handy and often cleans up your
measurements!
The shutter to the PMT on the PTI closes to protect it when the cover to the
instrument is open. However, it is easy to lean on the shutter that will cause it to open and
expose the PMT, leading to very noisy signal and potential damage to the PMT from
overexposure to light.
page 83
HPLC Protocol
For Chemical Biology Laboratory
By J. Frederick
(return to Contents)
Introduction
High performance liquid chromatography, commonly known as HPLC, has a variety of
applications in the chemical biology research laboratory. This protocol provides some
basic background theory, some tips for getting ready to use the HPLC for your particular
purification, and guidelines for doing an HPLC purification using our (fill in brand name)
instrument. Click here to refer to the HPLC glossary for definitions of many of the terms
used in the text.
There are many types of HPLC columns developed for specific applications. The right
choice of column is critical for obtaining good HPLC results. Column choice is governed
by characteristics of components in the mixture we wish to separate. For example, we can
separate components based on size, charge, hydrophobicity, aromatic character, even
chirality. Variable factors include the polarity of the stationary phase, column
dimensions, and pore sizes (which can be varied to allow certain sized analytes to pass
through at different rates). Another variable that impacts the efficiency of the HPLC
separation is the polarity of the mobile phase. Multisolvent delivery systems change the
page 84
polarity of the mobile phase over the course of an HPLC run, at a rate that defines the
"gradient" (e.g., 20% Buffer B to 100% Buffer B over 60 minutes). The use of a gradient
improves the separation of analyte mixtures of varying polarities.
The mechanics of the HPLC system are controlled by Windows-based software on a PC.
This software controls the gradient of the mobile phase, the solvent flow rate, mobile
phase pressure, and measures the signals produced by the detector. A specific HPLC
protocol is stored as a method, the parameters of which can be adjusted as necessary.
Finally, the results of your sample run can then be interpreted and printed in a variety of
report formats.
page 85
1. Very Important Note: Everything that goes into the HPLC must be filtered first,
through a 0.45 mm or 0.2 mm filter and special glassware to remove particles that
can get caught up on the column and interfere with absorption and separation. This
includes your buffers and your sample. Omission of this step can result in damage to
the instrument.
2. Sample preparation
The crude peptide, prepared by manual or automated synthesis, will be supplied as a
lyophilized (dried by freezing in a high vacuum) substance. For b-peptides, the sample is
dissolved in 50% H20/50% CH3CN (or a range of others; solvent selection depends on
solubility of the sample). Filter your sample.
3. Buffer Preparation
Buffer A and Buffer B are prepared according to the following recipes:
Buffer A Buffer B
80 mL CH3CN 3200 mL CH3CN
3920 mL H20 800 mL H20
2.4 mL TFA* 2.0 mL TFA*
*Safety precaution: Trifluoroacetic acid (TFA) is highly corrosive and causes severe
burns when inhaled or upon contact with skin. This chemical should only be handled in
the fume hood while wearing safety goggles, gloves, and protective clothing.
Filter your buffers, using the designated glassware and following the specific instructions
provided by your TA. This can be done prior to use and buffers stored at room
temperature until you are ready to use the HPLC.
4. HPLC Operation
Your TA will provide specific instructions pertaining to the use of the HPLC. Typically,
a run starts by attaching your buffers and washing the column (100% Buffer B for 5-10
minutes). Next allow the column to re-equilibrate to conditions that will start your run.
For a run with a gradient of 20% Buffer B to 100% Buffer B, this means allowing about
5-10 minutes for the starting conditions for injection to be achieved (that is, to get the
entire column in 20% Buffer A).
Once a specific separation method is specified, you may review the parameters such as
pump flow gradient, run time, and the PDA setup (acquisition). On some instruments,
you will need to specify the lamp used for detection. Your TA will supply the details for
the instrument you are using.
When making an injection, choose the amount based on the type of column you are using
and the approximate amount of your sample. For a- and b-peptides, the following general
guidelines apply:
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Column scale Amount peptide per injection
Analytical Up to 0.01 mg
Semi-preparative Up to 0.05 - 0.1 mg
Preparative Up to 0.1 – 0.5 mg
Use either a glass syringe or a disposable plastic syringe fitted with a luer lock needle
(only use flat-tipped needles). Before drawing up your sample, wash out the syringe
several times with Buffer B. Draw your sample into the syringe, then carefully remove
ALL bubbles from the sample by inverting the syringe, tapping gently, and expelling air
until liquid just appears at the needle tip. Load your sample as instructed by your TA.
You will want to adjust the view on the PC screen for convenient monitoring of the run,
which means selecting the appropriate wavelength(s). For a- and b-peptides, 214 nm (the
absorption frequency of peptide bonds) and 280 nm (the absorption of tyrosine and
tryptophan) are recommended. Notice the retention times listed (in minutes) at the bottom
of the graphs as well as in the status bar at the top of the screen (this may vary depending
on the software used; your TA will clarify this). You will need to record the retention
times as you collect peaks so you can correlate your fractions with peaks on the
chromatogram.
For the first injection of a peptide you’ve never purified before, you will need to carefully
analyze the output. To do this, label a set of 15-20 tubes (15-ml conical vials usually
work; you may want to do this ahead of time and loosen the caps so they are ready for
collecting peaks as they come off the column. Once you have collected all the relevant
peaks from the first injection, you will analyze them by mass spectrometry and determine
which fraction or fractions contain your peptide by looking for its molecular weight
(calculated in advance). Matching these fractions to their corresponding peaks will give
you the retention time for your molecule. At this point, further injections will be
simplified as you can accurately predict the retention time of your sample, and you’ll
know where to expect the peak containing your molecule.
The first peaks that come off the column (after 3-4 minutes dead time for the semi-prep
column, 5 minutes dead time for a prep column) represent a variety of leftovers from the
synthesis (usually incomplete removal of reagents during wash steps). Once you get
beyond this point you should collect every peak as it comes off the column, noting the
retention time (for example: 11.23-11.5) for each numbered tube. Try to separate
shoulders from main peaks, and isolate peaks that appear within multiple peaks. Keep in
mind that the method you choose will impact the appearance of the chromatogram, and
hence your ability to collect a pure, isolated fraction. For example, a longer method will
give better resolution, but broader peaks. You will get better at this technique with
practice, and bear in mind that it usually takes at least two passes through a column to
purify a crude peptide synthesis.
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HPLC Glossary
For a comprehensive list of terms, see
http://www.waters.com/watersdivision/images/aboutus/hplcglossary.htm.
Column – A tube containing the stationary phase. The stationary phase differentially
interacts with the sample’s constituent compounds as they are carried along in the mobile
phase.
Fraction – A sample collected from the instrument after it has flowed through the
column and passed by the signal detector.
Gradient – The change in mobile phase composition over time. This can be continuous
or stepwise.
Mobile phase – The solvent that moves the mixture of compounds through the column.
Retention time – The elapsed time between injection of a sample and appearance of a
peak maximum.
Stationary phase – The immobile phase in the chromatographic process. In HPLC, this
is a solid material packed inside a column.
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Diagram 1
B + A represents a mixture of analytes to be separated. A moves faster through the
column than B, and will therefore have a shorter retention time. The small peak to the left
of A represents unwanted material such as degradation products or leftovers from
synthesis. This material often shows up as several peaks, sometimes larger than the
product peak(s), at an early point in the separation.
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I. Introduction
Mass spectrometry is a powerful tool used for studying the masses of atoms,
molecular fragments, and molecules. In general, molecules in the gas phase (or
species desorbed from a condensed phase) are ionized, and the ions are then
accelerated by an electric field and separated on the basis of their mass-to-charge
ratio (m/z). For an ion with a charge of +1, m/z will be numerically equal to the mass.
The electron ionization that converts molecules to ions can not only remove electrons,
but may impart so much energy that the molecule fragments. The molecular ions are
deflected by a magnet as they travel through the analyzer tube toward the detector. A
mass spectrum is a chromatogram presenting the signal intensity (y-axis) versus m/z
(x-axis). See Figure 1 below. The peak intensities are expressed as a percentage of the
most intense signal (the base beak). A time-of-flight (TOF) mass spectrometer
separates ions with identical kinetic energy but different m/z, since lighter ions travel
faster than heavier ones. Interpretation of the fragmentation patterns and isotopic
peaks can provide valuable clues for the structure determination of organic molecules.
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Figure 1. Mass Spectrum of bovine serum albumin obtained using MALDI-TOF. Figure
adapted from the American Society for Mass Spectrometry website
(http://www.asms.org/whatisms/p12.html).
Selecting a matrix
Selection of proper matrix is important for getting a good mass spec, since the matrix
plays a key role in ionization. The chart below suggests guidelines to consider.
Matrix Application
Sinapinic Acid Peptides and proteins greater than 10 kDa
(3,5-dimethoxy-4-hydroxy cinnamic acid) in mass
CHCA Peptides and proteins less than 10 kDa in
(a-cyano-4-hydroxycinnamic acid) mass
THAP Small oligonucleotides less than 3.5 kDa in
(2,4,6-Trihydroxyacetophenone) mass
HPA Large nucleotides greater than 3.5 kDa in
(3-hydroxypicolinic acid) in diammonium mass
citrate
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I have found that aPP derived molecules and other hydrophobic peptides work well in
sinapinic acid. For smaller peptides (18-24 mer), CHCA can be used.
CHCA:
In an eppendorf tube, weigh out 10 mg of CHCA. Add 400 µL of deionized water, 100
µL of 3% TFA and 500 µL of acetonitrile to the matrix. Vortex for 1 minute to disslove,
then centrifuge for 1 minute to precipitate any undissolved sinapinic acid. Use only the
supernatant for applications.
Note 1: Buffers, salts and detergent retard the ionization of the matrix. As far as possible,
avoid using the last two (salts and detergents). If you must use a sample prepared in a
buffer, increase the concentration of TFA in the matrix stock to enhance sample
ionization.
Note 2: If the dry matrix is a mustard-yellow color instead of bright yellow, it may
contain impurities. To purify, dissolve CHCA in warm ethanol. Filter and add 2 volumes
of deionized water. Let the solution stand in the refrigerator for 2 hrs. Filter and wash the
precipitate with cold water.
THAP
Make a 50 mg/mL solution of diammonium citrate in deionized water. Dissolve 10 mg of
THAP in 50% acetonitrile/deionized water. Combine 8:1:: THAP solution:diammonium
citrate solution.
Note: For oligonucleotide applications, do not use HPLC grade water for sample
preparation. Use deionized water only.
HPA:
Make a 50 mg/mL solution of diammonium citrate in deionized water. Dissolve 50 mg of
THAP in 50% acetonitrile/deionized water. Combine 8:1:: THAP solution:diammonium
citrate solution.
Note: For oligonucleotide applications, do not use HPLC grade water for sample
preparation. Use deionized water only.
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Peptides and proteins: 1-100 pmol/µL (Lower concentration for smaller peptides and
higher concentration for proteins and larger peptides, i.e. > 5 kDa)
Oligonucleotides: 10-100 pmol/L
Samples should be preferably dissolved in water. If insoluble in water, add acetonitrile to
the solution (up to 50%) and then 0.1% TFA to increase solubility. Consult your TA for
specific guidelines on sample stock solutions.
III. Troubleshooting:
If you do not see any signal on the mass spec, it could be due to the following reasons:
Be sure to avoid:
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1) Using organic solvents to dissolve samples - causes the sample to spread out and not
crystallize properly.
2) Touching the surface of the sample plate with the pipet tip - causes uneven
crystallization.
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I. Experimental Strategy
The experimental strategy for panning of phage particles against DNA targets is
outlined in the figure below. The phage particles are generated from the library in XL1-
Blue cells upon infection with M13KO7 helper phage. The target DNA (for example:
hsCRE, ATGAC) is biotinylated on one strand and can be immobilized using
streptavidin-coated magnetic beads. The phage particles are then exposed to the DNA.
Washing and elution of the beads isolates only the desired phage particles. These are then
reinfected into XL1-Blue cells, serially diluted, and plated to determine the number of
retained phage. For comparison, a sample of the phage particles without exposure to
DNA are infected into XL1-Blue cells, serially diluted, and plated to determine the input
titer. From the number of colonies, the percent retention of the phage particles on the
DNA target site is determined. To isolate the phage particles with the highest specificity
and binding, multiple rounds of selection will be necessary.
For a single round of panning, this protocol will require five days. In the first and
second days, all of the necessary solutions are prepared, the beads are washed and
blocked, and the cell cultures are infected to produce phage particles. The following day,
the biotinylated DNA is added to the beads, and the phage particles are isolated. The
phage particles are mixed with the beads, then immobilized on a magnet. Washing and
elution allows for the selection of high affinity binding phage particles. These selected
phage particles are then reinfected into XL1-Blue cells and plated. On the final day, the
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plated cells are tallied to determine the retention percentages.The outlined procedure is
effective when performing 4 assays. Typically, two of these can be positive and negative
controls. For example: pCANTAB-007 and pCANTAB-APP (or beads containing no
DNA).
II. Solutions
1M NaOH 2M MgCl2
4.0 g NaOH (40 g/mol) 40.66 g MgCl2•H20 (203.30 g/mol)
Add H2O to 100 mL Add H2O to 100 mL and autoclave
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III. Protocol
– Day 1 –
Assays
• _pCANTAB-negative control • _pCANTAB-Library A
• _pCANTAB-positive control • _pCANTAB-Library B
Assay Starter Cultures
• 6:45pm - Add -control and +control glycerol stabs to 5 mL 2X YT-AG. Add
Library A and Library B glycerol stocks (not pool 0) to 10 mL of 2X YT-AG in
50 mL orange cap tubes. Streak XL1-blue glycerol stock on LB-tet plate. Incubate
at 37ºC.
– Day 2 –
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– Day 3 –
Isolation of Phage
• 9:45am - Spin the phage producing cell cultures (10 mL) at 2,500 rpm for 20
minutes. Get ice. Thaw dI-dC (1 µg/µL) and 1.0µM duplex DNA.
• Filter the broth through a 0.45 µm filter using a 10 mL syringe into a sterile
centrifuge tube. (The cells may be discarded.)
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centrifuge and rotor SA-600 or the Beckman centrifuge and rotor JA-20 to 4°C.
• 11:05am - Spin the broth/PEG solutions at 20,000 G, (SA-600-11,800 rpm, JA-20-
13,000 rpm) for 30 minutes.
• After centrifugation, decant the broth into bleach, then invert the tubes on paper
towels to dry the phage particle pellet. (Translucent white precipitate may not be
visible.)
• A "phage solution" for each assay is prepared by adding 1 mL of Buffer A* to each
centrifuge tube containing a phage pellet (after drying). Chill on ice.
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XL1-blue Culture
• 4:30pm - Don't forget about the XL1-blue cells from the starter culture! After
growing to an OD600 of 0.8, aliquot 7 mL into Falcon culture tubes (2 x number
of assays 8).
Serial Dilutions
• Make multiple serial dilutions for the elution titers by adding 100 µL of the 7 mL
culture to 900 µL of 2X YT-AG (101, 102, 103, 104, 105, & 106 dilutions).
• Likewise, make multiple serial dilutions for the input titers by adding 100 µL of
culture to 900 µL of 2X YT-AG (101, 102, 103, 104, 105, 106, 107, 108, & 109
dilutions).
• Plate a 20 µL droplet from each of the above serial dilutions on SOB-AG agar plate
(4 per plate) and incubate overnight at 37°C.
Glycerol Stocks
• Glycerol stocks of XL1-blue cells containing selected Libraries A and B phagemids
are prepared from elution titers by adding 0.8 mL to 0.4 mL 50% glycerol and
freezing.
– Day 4 –
– Day 5 –
Notes
• Timepoints are approximate.
• Discard into bleach anything that comes into contact with phage. Separate
pipetmen for phage may be used. Filter tips must be used.
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• The stringency of the selections can be varied by changing the temperature, number
of washes, length of washes, or by adding competitor DNA.
1 mL phage
7 mL culture 0.4 mL eluted 1
Elution # soln
= 10x
Titer colonies 0.02 mL 0.4 mL bind 10 mL
0.2 mL infect
plated mix culture
1 mL phage
7 mL culture 1
Input # soln
Titer
=
colonies
10x
0.02 mL
10 mL
0.1 mL infect
plated culture
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Elution
Negative
control
Input
Elution
Positive
control
Input
Elution
Library A
Input
Elution
Library B
Input
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Required solutions
TBST
Dissolve 10 mL of 1M Tris.HCl (pH 8.0)
8.7 g NaCl
0.5 mL Tween-20
in 1L of diH20. Sterile filter.
Protocol
– Day 1 –
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– Day 2 –
Add Phage:
• Start a 10 mL 2xYT-AG growth for each clone in 50 mL orange cap centrifuge tubes
from 1mL of overnight culture. Grow at 37 °C until log phase.
• Add 400 µL M13K07 helper phage to each.
• Shake at 37 °C for 1 hr.
• Spin cells down at 2500 rpm for 20 min in delicase centrifuge.
• Resuspend cells in 10 mL 2xYT-AK.
• Grow phage overnight (12 hours).
– Day 3 –
Phage precipitation:
Spin cells down at 2500 rpm for 20 min in delicase centrifuge.
Discard cells, filter broth through 0.45 µm filter into centrifuge tube.
Add 2 mL 0.2 µm filtered PEG/NaCl to each tube.
Incubate on ice for 45 min.
Centrifuge cells for 35 min at 13000 rpm at 4 °C.
Discard supernatant; dry pellets for ~ 2 min.
Resuspend pellet in 1 mL TBST buffer.
Panning:
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Infection:
Infect 5 mL XL1Blue culture with 100 µL input and 5 mL with 100 µL output phage.
Incubate at 37 °C for 1 hr.
Titering:
Make neat - 107 serial dilutions of output phage.
Make 101 - 1010 serial dilutions of input phage.
Plate 20 µL of dilutions of output phage and input phage on SOBAG plates.
Incubate at 30 °C overnight (16 hours).
– Day 4 –
Pick 20 colonies from output titer plates and grow in 3 mL 2xYT overnight at 37˚C.
– Day 5 –
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Amino acid analysis is a technique used to characterize a protein’s amino acid content
and the concentration of a given sample. We out-source this service through the Keck
facility (formally the HHMI Biopolymer – Keck Foundation Biotechnology Resource
Foundation) at Yale. This facility is located in the Boyer Center of the Medical School
and can be accessed online (http://info.med.yale.edu/wmkeck/prochem/aaa.htm).
The Keck website provides detailed information about what happens to your protein
during the amino analysis process. In brief, the protein is completely hydrolyzed and then
subjected to chromatographic analysis (HPLC) against amino acid standards.
“Amino acid analysis is carried out on a Beckman Model 7300 ion-exchange instrument
following a 16 hr hydrolysis at 115 degrees C in 100 µl of 6 N HCl, 0.2% phenol that
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also contains 2 nmol norleucine. The latter serves as an internal standard to correct for
losses that may occur during sample transfers, drying etc. After hydrolysis, the HCl is
dried in a Speedvac and the resulting amino acids dissolved in 100 µl Beckman sample
buffer that contains 2 nmol homoserine with the latter acting as a second internal standard
to independently monitor transfer of the sample onto the analyzer. The instrument is
calibrated with a 2 nmol mixture of amino acids and it is operated via the manufacturer's
programs and with the use of their buffers. Data analysis is carried out on an external
computer using Perkin Elmer/Nelson data acquisition software.
During acid hydrolysis asparagine will be converted to aspartic acid and glutamine to
glutamic acid. During the HPLC analysis that follows, cysteine co-elutes with proline;
and methionine sulfoxide, which is a common oxidation product found in
peptides/proteins, co-elutes with aspartic acid. Hence, following normal acid hydrolysis,
glutamine and asparagine are not individually quantified and it is possible that the
methionine value will be low and (generally to a lesser extent) that the aspartic acid and
proline values will be somewhat high. Improved quantitation of cysteine and methionine
can be obtained by requesting prior oxidation with performic acid, which converts both
methionine and methionine sulfoxide to methionine sulfone and cysteine and cystine to
cysteic acid. Generally, however, performic acid oxidation destroys tyrosine. Best
quantitation of tryptophan is generally obtained by requesting hydrolysis with
methanesulfonic acid (MSA) instead of hydrochloric acid. The procedure used in this
instance is to carry out the hydrolysis with 20 µl MSA for 16 hrs at 115C. After
hydrolysis, the sample is neutralized with approximately 200 µl 0.35 M NaOH and 100
µl (50% of the sample) is then analyzed on the Beckman 7300. Please keep in mind that
since we believe the overall extent of hydrolysis with MSA is less than with HCl, we do
not recommend MSA hydrolysis for use in quantifying the concentration of protein stock
solutions.”
Your TA will provide detailed information about how to prepare your sample for
submission, and how to fill out the form that must accompany each submission. The
forms can be downloaded from http://keck.med.yale.edu/yaleforms.htm.
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The principle behind this method is based upon the mathematical description of how a
particle behaves when subjected to a centrifugal force. The sedimentation velocity (how
fast a particle moves toward the bottom of a tube) depends on factors including the mass,
shape and density of the particle.
Gradient centrifugation is used for the separation of proteins with different sedimentation
coefficients. A linear density gradient is formed by mixing high and low density
solutions in a centrifuge tube. A solution containing the proteins to be separated is then
layered on the top. As the rotor is spun, the proteins move through the solution to
separate at rates dependent upon their sedimentation coefficients. The separated bands of
protein can be collected for analysis by piercing a hole through the bottom or side of the
tube and carefully withdrawing drops of solution.
Sedimentation equilibrium is used for the direct determination of the mass of a protein.
Samples are centrifuged at low speeds to counterbalance sedimentation with diffusion.
This method of mass determination is highly accurate and can be used under non-
dentaturing conditions to preserve the native quaternary structure of multimeric proteins.
By comparing the estimated mass of denatured polypeptide chains elicited from SDS-
polyacrylamide gel electrophoresis to the mass of the intact protein determined by
sedimentation equilibrium analysis, one can ascertain how many copies of each
polypeptide chain are in the protein. This method can be used to determine whether a
peptide exists as a monomer, or whether it dimerizes or forms higher aggregates in
solution.
The Schepartz laboratory has established a collaboration with Jim Lear at the University
of Pennsylvania for analytical centrifugation analysis. Your TA will provide specific
information about preparing your particular samples for submission.
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Operation
The lyophilizer is maintained by the lyophilizer Czar. Other users should not need
to worry about defrosting the system or changing pump oil. The RC5 Hybrid vacuum
pump on this system is very durable and will not need a regular oil change. But the pump
should NOT be shut down, as this defeats the mechanism by which the pump continually
clears itself of 'inhaled' organics and water. The lyophilizer Czar will shut the pump off
for short times while defrosting the drying chamber (without allowing the oil to cool
significantly), but any other need to shut the pump off should be cleared by the Czar.
To add a sample to the system, first confirm that the temperature is below -40˚C
and the pressure is AT LEAST registering a value on the control panel. Ideally, samples
should only be added when the green indicator light is blinking or solid. In situations
where many samples have been added to the system, this may be impractical, as the
vacuum in the system may not be capable of getting this low. In these cases, as long as
the vacuum has stabilized, it should be okay to add your sample. If the control panel does
not show a numerical pressure (i.e., 100 x10-3 mbar), but instead reads "HI", do not add
samples, as the vacuum is not sufficient for lyophilization.
Add your sample (pre-frozen on dry ice) by connecting either your flask or the
Labconco container containing your vials to the chamber with the appropriate fittings. To
open your sample to the vacuum system, turn the grey knob slowly 180˚. The 'vent'
position (flask closed to system) is when the flat portion of the grey knob is lined up with
the hole in the black seal. Turning the grey knob 180˚ opens the flask to the vacuum
chamber. At this point any residual liquid in your sample may bubble or otherwise cause
your sample to shift in your flask. After you have opened the flask to the vacuum
chamber, observe the sample a few moments to ensure that any shifting or bubbling does
not upset your sample too much, and to make sure your sample stays solid initially.
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To remove your sample from the lyophilizer, reverse the process outlined above
by turning the grey knob 180˚ to the ‘vent’ position. Beware that turning the grey knob to
the vent position (flat surface lined up with the hole in the seal) will allow room air into
your sample flask. This flask is under high vacuum, and the air will rush in very
vigorously. Take care to turn the knob slowly, or your dry sample will blow around in
your flask, possibly flying out of your flask. Many a sample has been violently blown
into the drying chamber in this way.
Problems/Fixes
For efficient lyophilization, your sample must be frozen. Unlike the Speed-Vac,
the lyophilizer will not reduce liquid samples in a desirable way. If you attempt to dry a
wet sample, you will learn why the Speed-Vac uses a centrifuge system (and you will
never again take the fact that your sample stays inside your flask for granted).
In cases where you use a round bottom flask directly attached to the system, it is
normal for frost to form on the outside of the flask. This will also occur with the
Labconco glassware if the contents (vials, etc.) are in contact with the outer glass
container. Keep in mind that this frost melts, so you may want to place a paper towel
under the flask to absorb the water.
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The PhastSystem
by Kevin Rice
adapted for Chemical Biology Laboratory by J. Frederick
(return to Contents)
General Considerations
The PhastSystem is designed for quick, easy, and reproducible electrophoresis
applications for both protein and nucleic acid samples. The system includes pre-cast
polyacrylamide gels and buffers in a variety of flavors to accommodate many of your
electrophoretic needs. While Phast gels can be invaluable for rapid qualitative analysis,
using such small gels does introduce limitations relative to hand-poured gels. First, the
resolution of many molecules within a very narrow molecular weight range (DNA
sequencing for example) usually requires a longer separation zone. In addition, these very
thin gels are inadequate for preparative electrophoresis. Finally, only polyacrylamide gels
can be used, precluding the efficient separation of large (>1000 bp) DNA molecules.
Despite these limitations, Phast gels are an attractive option for any of the following
applications frequently encountered in the Schepartz laboratory:
Other applications which are not as common in this laboratory and therefore
not discussed here are: isoelectric focusing, 2-dimentional protein electrophoresis,
transfer for western blotting, and native protein electrophoresis (for more
information about applications, see the Amersham Biosciences website
http://www1.amershambiosciences.com/aptrix/upp01077.nsf/Content/Products?Ope
nDocument&parentid=40314&moduleid=40316).
The PhastSystem also includes a development chamber most useful for silver-
staining gels. Both protein and nucleic acid gels can be silver-stained. Phast gels can also
be stained using standard protocols with Coomassie Brilliant Blue, ethidium bromide,
and other stains.
Described here is the general procedure for operating the PhastSystem apparatus.
Attached is the list of current Phast programs and silver-stain solutions I will not describe
method programming here. Should this be necessary, the PhastSystem literature describes
that procedure.
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soaked into 3% agarose strips that are positioned at either end of the gel. These strips
contain either native or denaturing (with SDS) buffers that subsequently enter the gel
matrix giving the gel the desired characteristics. Buffer strips can be regenerated by
successive soaks in the appropriate buffer. In addition, strips can be soaked in different
buffers to generate buffer environments not provided by Pharmacia. The best
combination of Phast gel and buffer strip is not necessarily obvious. Often, one must try
different procedures until satisfactory electrophoretic separation is obtained. Much
information is available in both the Pharmacia catalog and their website. What you need
to get started is provided here.
Sample Preparation
The load volume on a Phast gel can be no more than 4 µL. In that case, the comb
for one gel has 6 wells. We also have 8-well combs, but they hold only 1 µL per well.
Keep this in mind when preparing your samples, as well as what staining procedure you
plan to use. Coomassie stain detects most protein bands at 50-100 ng, while silver stain
detects as little as 0.3 ng per band for protein and 20 pg per band for DNA. Remember to
keep these facts in mind when preparing marker/ladder samples. Use the same amount of
loading buffer (with glycerol or ficoll and dyes) as you would on any other gel. Always
prepare at least twice needed volume to allow for slips-of-hand (which happen more
often here than with other kinds of gels).
Apparatus Setup
Chosen gel(s), which are stored in the refrigerator, are removed from their
packages by cutting along the dotted lines. If you fail to cut along the lines, you will be
demoted back to kindergarten. The front of the package corresponds to the top of the gel.
The gel itself is affixed to a piece of plastic. The top of the gel is protected by a thinner
piece of plastic that must be carefully removed before use. Often, this protective layer
will stick to the package when peeled away from the gel. Once removed, be careful not to
scratch the gel. If this happens, discard the gel and get another one as a scratch in the gel
will in all likelihood ruin electrophoresis. You can bend back the trapezoidal nub to aid
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later handling of the gel. Lift up all hinged parts in the PhastSystem separation unit and
align each gel with the red outline on the white surface (gel side up). If you are using
only one gel, it does not matter which position you choose. Put the chosen buffer strips in
the appropriate positions on the clear-plastic removable buffer strip holder, and put the
holder on top of the gels, putting the pins in the back of the surface through the holes in
the holder. Press down on the strips gently to ensure they make contact with the gel.
Lower the hinged parts. Again, press on the electrodes that touch the buffer strips to
ensure contact.
Turn the instrument on (button in the back). Press the “SEP temp stand by” button
on the console. This will allow you to equilibrate the temperature to 15°, at which most
programs run their gels. The readout should give the current temperature as well as the
set temp of 15°. If the “(OFF)” is seen, press the “do” button such that “(ON)” shows up
on the display.
Running a Program
Once loaded and ready to go, press “SEP start stop.” Enter the number of gels (1
or 2) and press “do.” Enter the program number (the different programs are on the sheet
attached here and posted above the PhastSystem) and press “do.” The program will now
run automatically. It will not stop until you stop it. Track the progress of the marker dyes
as you would for a hand-poured gel and stop it when the gel has sufficiently run. Stop by
pressing “SEP start stop” and then “do” to verify the stopping.
Staining
If you are staining your gel with Coomassie or ethidium bromide, treat it as you
would any other gel. If you plan to use the development chamber for silver staining, place
your gels in the wire brackets within the chamber. Make sure all silver stain solution
bottles are full (the locations and recipes for the solutions are in the attached sheet and
above the PhastSystem). Staining 1 or 2 gels involves no differences in staining protocol.
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Press “DEV start stop” and then the number of the development program and then “do.”
The gel will then be stained automatically.
Apparatus Shutdown
Wipe all used components with a wet Kimwipe and then a dry Kimwipe. Clean up
the bench area around the PhastSystem or Stacey will hurt you. Shut off apparatus when
completed.
Silver Solutions
1 20% TCA (trichlororacetic acid) (TCA solid in Room 114 fridge)
2 50% Ethanol / 10% HOAc
3 10% Ethanol / 5% HOAc (stock bottle on bottom shelf)
4 5% Gluteraldehyde (glutaric dialdehyde) (stock in flammables cabinet)
5 dH2O
6 0.4% Silver nitrate (in near fridge)
7 Developer (make fresh)
8 Background Reducer (make fresh)
9 5-10% Glycerol
Developer:
1 mL 2% formaldehyde (in near fridge)
150 mL 2.5% Na2CO3 (stock bottle on bottom shelf)
2% Formaldehyde:
1 mL 37% formaldehyde (in flammables cabinet)
17 mL dH2O
Background Reducer:
3.7 g Tris-HCl (near balance in hood)
2.5 g sodium thiosulfate (near balance in hood)
100 mL H2O
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For additional information about the Phastsystem, the user manual is available online as a
pdf file:
http://www1.amershambiosciences.com/aptrix/upp00919.nsf/(FileDownload)?OpenAgent
&docid=69BE4F0F5FE010BBC1256AB100084A40&file=80132015.pdf
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Theory
The speed-vac is used to concentrate small-volume samples. Under vacuum (very
low pressure), the vapor-liquid equilibrium of the solvent is shifted towards the gas
phase, while your sample (DNA, peptide, etc.) remains primarily in the solid phase.
Therefore, using a vacuum you can easily remove solvent with very little stress on your
solute, leaving you with a dry, solid sample (plus salts that were present in the solvent
buffer, etc.). You can then resuspend the sample in the desired amount of any buffer you
want. For larger volumes, the lyophilizer is used to freeze-dry samples.
2. Before opening the lid to the speed-vac, you must release the vacuum. This is done
by turning the bleed valve perpendicular to the line (closed position).
5. When concentrating very hot samples (i.e. freshly end-labeled DNA), the samples
MUST be frozen and in a screw cap vial with the cap on.
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6. Sign your name, time, and sample type on the log beside the speed-vac. Also indicate
if you do not want your samples exposed to heat or light.
7. To remove samples follow the same procedure for releasing the vacuum and opening
the sample chamber. Remember to turn the speed-vac back on after you remove your
samples.
8. If taking unlabeled samples out of the radioactive (hot) speed-vac (i.e. basic samples)
make sure that they are not radioactive (by checking with a Geiger counter) and be
aware that other samples may be hot. If you remove samples from the hot speed-vac
always check your gloves to make sure the inside of the speed-vac is not hot.
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Ultraviolet-Visible Spectroscopy
by Alain Martelli
adapted for Chemical Biology Laboratory by J. Frederick
(return to Contents)
I. Theoretical principles
Introduction
Many molecules absorb ultraviolet (UV) or visible light. The absorbance of a
solution increases as attenuation of the beam increases. Absorbance is directly
proportional to the path length, b, and the concentration, c, of the absorbing species,
according to the Beer-Lambert Law (see below):
A = ε bc
where ε is a constant of proportionality called the molar absorptivity. Different molecules
absorb radiation of different wavelengths. An absorption spectrum will show a number of
absorption bands corresponding to structural groups within the molecule.
Electronic transitions
The absorption of UV or visible radiation corresponds to the excitation of outer
electrons. There are three types of electronic transitions to be considered:
1. Transitions involving #, s and n electrons.
2. Transitions involving charge-transfer electrons
3. Transitions involving d and f electrons
When an atom or molecule absorbs energy, electrons are promoted from their
ground state to an excited state. In a molecule, the atoms can rotate and vibrate with
respect to each other. These vibrations and rotations also have discrete energy levels,
which can be considered as being packed on top of each electronic level.
Absorption of UV and visible radiation in organic molecules is restricted to
certain functional groups (chromophores) that contain valence electrons of low excitation
energy. The spectrum of a molecule containing these chromophores is complex, because
the superposition of rotational and vibrational transitions with the electronic transitions
gives a jumble of overlapping lines that appears as a continuous absorption band.
Charge-transfer absorption
Many inorganic species show charge-transfer absorption; these are called charge-
transfer complexes. For a complex to demonstrate charge-transfer behavior, one of its
electrons must be able to be donated, and another component must be able to accept the
electron. Absorption of radiation then involves the transfer of an electron from the donor
to an orbital associated with the acceptor. Molar absorptivities from charge-transfer
absorption are large (greater than 10,000 L.mol-1.cm-1). Depending on the complex,
charge-transfer complexes can absorb almost anywhere in the UV-Vis range.
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s s* transitions:
An electron in a bonding s orbital can be excited to the corresponding antibonding
orbital, though the energy required for this is large. For example, methane (which has
only C-H bonds, and can only undergo s s* transitions) cannot be seen in typical UV-
Vis spectra (200 – 700 nm).
n s* transitions:
Saturated compounds containing atoms with lone pairs (non-bonding electrons)
are capable of n s* transitions. These transitions usually need less energy than s s*
transitions. They can initiated by light whose wavelength is in the range 150 – 250 nm.
The number of organic functional groups with n s* peaks in the UV region is small.
n #* and # #* transitions:
Most absorption spectroscopy of organic compounds is based on transitions of n
or # electrons to the #* excited state. This is because the absorption peaks for these
transitions fall in an experimentally convenient region of the spectrum (200 – 700 nm).
These transitions need an unsaturated group in the molecule to provide the # electrons.
Molar absorptivities from n #* transitions are relatively low, and range from 10
to 100 L.mol-1.cm-1. # #* transitions normally give molar absorptivities between 1,000
and 10,000 L.mol-1.cm-1.
The solvent in which the absorbing species is dissolved also has an effect on the
spectrum. Peaks resulting from n #* transitions are shifted to shorter wavelengths
(blue shift) with increasing solvent polarity. This arises from increased solvation of the
lone pair, which lowers the energy of the n orbital. Often (but not always), the reverse
(red shift) is seen for # #* transitions. This is caused by attractive dipole forces
between the solvent and the absorber, which lower the energy levels of both the excited
and unexcited states. This effect is greater for the excited state, and so the energy
difference between the excited and unexcited states is slightly reduced. This results in a
small red shift. This effect also influences n #* transitions, but is overshadowed by the
blue shift resulting from solvation of lone pairs.
The Beer-Lambert Law
The diagram below shows a beam of monochromatic radiation of radiant power
P0 directed at a sample solution. Absorption takes place and the beam of radiation leaving
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the sample has a radiant power P. The amount of radiation absorbed may be measured in
a number of ways:
Transmittance, T = P/P0
%Transmittance, %T = 100 T
Absorbance:
A = log10 (P0 / P)
= log10 (1 / T)
= log10 (100 / %T)
= 2 – log10 %T
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260 nm (subtracting the "blank" absorbance) and then simply calculating concentration
via a standard factor as per the Beer-Lambert law.
An absorption of 1.0 is equivalent to approximately:
50 µg/mL double-stranded DNA (dsDNA)
33 µg/mL single-stranded DNA (ssDNA)
40 µg/mL single-stranded RNA
30 µg/mL for ssDNA oligonucleotides.
For more precise calculation methods, see the separate section on DNA technical
information or use the biopolymer calculator on our website.
The purity of a nucleic acid sample can be assessed by calculating the ratio
between absorbances at 260 nm and 280 nm. This ratio (A260/A280) is used to estimate
purity because proteins absorb more strongly at 280 nm. Pure DNA should have a ratio of
approximately 1.8, whereas pure RNA should give a value of approximately 2.0.
Absorption at 230 nm reflects contamination of the sample by substances such as
carbohydrates, peptides, phenols or aromatic compounds. The ratio A260/A230 should be
approximately 2.2 for pure nucleic acid samples.
Materials:
1. Lyophilized bovine plasma gamma globulin or bovine serum albumin (BSA)
2. Coomasie Brilliant Blue 1
3. 0.15 M NaCl
4. Spectrophotometer and tubes
5. Micropipettes
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1. Prepare a series of protein standards using BSA diluted with 0.15 M NaCl to final
concentrations of 0 (blank, NaCl only), 250, 500, 750 and 1500 µg BSA/mL. Also
prepare serial dilutions of the unknown sample to me measured.
2. Add 100 µL of each of the above to a separate test tube
3. Add 5.0 mL of Coomasie Blue to each tube and mix by vortex or inversion.
4. Adjust the spectrophotometer to a wavelength of 595 nm, and record the blank using
the tube from step 3 which contains no BSA.
5. Wait 5 minutes and read each of the standards and each of the samples at 595 nm
wavelength.
6. Plot the absorbance of the standards versus their concentration. Compute the
extinction coefficient and calculate the concentrations of the unknown samples.
Materials:
1. 0.15% (w/v) sodium deoxycholate
2. 72% (w/v) trichloroacetic acid (TCA)
3. Copper tartrate/carbonate (CTC)
4. 20% (v/v) Folin-Ciocalteu reagent
5. Bovine Serum Albumin (BSA)
6. Spectrophotometer and tubes
7. Micropipettes
Procedure:
1. Prepare standard dilutions of BSA of 25, 50, 75 and 100 µg/mL. Prepare appropriate
serial dilutions of the sample to be measured.
2. Place 1.0 mL of each of the above into separate tubes. Add 100 µL of sodium
deoxycholate to each tube.
3. Wait 10 minutes and add 100 µL of TCA to each tube.
4. Centrifuge each tube for 15 minutes at 3,000 G and discard the supernatant.
5. Add 1.0 mL of water to each tube to dissolve the pellet. Add 1.0 mL of water to a
new tube to be used as a blank.
6. Add 1.0 mL of CTC to each tube (including the blank), vortex and allow to set for 10
minutes.
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7. Add 500 µL Folin-Ciocalteu to each tube (including the blank), vortex and allow to
set for 30 minutes.
8. Turn on and zero the spectrophotometer to a wavelength of 750 nm. Use the blank
from Step 7 to adjust for 100% T.
9. Read each of the standards and samples at 750 nm.
10. Plot the absorbance of the standards versus their concentration. Compute the
extinction coefficient and calculate the concentrations of the unknown samples.
Notes
The Lowry method depends on the presence of tyrosine within the protein to be
measured. The standard protein must contain approximately the same number of tyrosine
residues as the sample, or the procedure will be inaccurate. If there are no tyrosine
residues in the sample to be measured, the Lowry method of protein determination is
useless and you should try the Bradford assay instead. In general, the Bradford assay is
the method of choice for protein determinations.
Materials:
1. Biuret Reagent
2. Bovine serum albumin (BSA)
3. Spectrophotometer and tubes
Procedure:
• Prepare standard dilutions of BSA containing 1, 2.5, 5.0, 7.5 and 10 mg/mL. Prepare
serial dilutions of the unknown samples.
• Add 1.0 mL of each of the standards, each sample, and 1.0 mL of distilled water to
separate tubes. Add 4.0 mL of Biuret reagent to each tube. Mix by vortexing.
• Incubate all of the tubes at 37 ºC for 20 minutes.
• Turn on and adjust a spectrophotometer to read at a wavelength of 540 nm.
• Cool the tubes from Step 3, blank the spectrophotometer and read all of the standards
and samples at 540 nm.
• Plot the absorbance of the standards versus their concentration. Compute the
extinction coefficient and calculate the concentrations of the unknown samples.
Notes:
The Biuret reaction was one of the first for the determination of protein
concentration. It remains as a rapid determination, but is not very accurate. It is useful
during protein separation procedures since there are fewer salt interference reactions than
with the Bradford or Lowry techniques. The color formed is stable for only 1 or 2 hours
and consequently all spectrophotometer readings must be made as soon as possible after
the incubation step.
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of competent cells, which must be in a specific phase of growth, or for inducing protein
expression in a bacterial culture.
There are two UV-Vis spectrophotometers available for use, a departmental one
in the instrument center and one housed in the Schepartz lab. Your TA will supervise
your operation of the instrument until you are comfortable with the technique. Using a
UV-Vis is quite simple, but there are a few important things to keep in mind to protect
the machine and ensure high quality data.
There are designated cuvettes for use by Chemical Biology Laboratory students.
For UV absorbance, use the quartz cuvette (volume = 100 mL). Be extremely careful
because they are fragile and expensive! Wash them with distilled water and then ethanol,
and use Kimwipes if you need to wipe them. If they are very dirty, they may need a bath
in concentrated acid (hydrochloric acid or nitric acid); consult your TA for assistance.
You should always clean a cuvette before and after each use. For bacterial cell densities
(OD600), you can use disposable cuvettes (volume = 3 mL).
For the practical use of the Schepartz B640 spectrophotometer, you can refer to
the useful QUICK REFERENCE provided in the operating instruction manual (first page
in the manual). You will find all the information you need to run the different options
(Rediread, Rediscan, Fixed wavelength, Wavelength scan, Time drive, Graphic
manipulations).
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Solvent UV cutoffs
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Western Blots
by Tanya Schneider
adapted for Chemical Biology Laboratory by J. Frederick
(return to Contents)
Western blotting is useful in detecting a protein of interest that may be mixed with
others (such as in a cell lysate) or verifying the identity of a protein on a gel. In general, a
mixture of proteins is resolved using a denaturing acrylamide gel. The separated proteins
are then transferred from the gel onto a nitrocellulose or PVDF membrane. The protein of
interest is probed by incubating the membrane with a specific antibody. The membrane
next is incubated with a secondary antibody that recognizes any bound primary antibody.
The secondary antibody generally enables (through a variety of mechanisms) eventual
identification of any band on the membrane that was bound by the antibody. This
technique is limited by the success of the gel separation, the transfer step and the
specificity of the antibody.
Some people have success using the Phast system to run and transfer a gel to a
membrane. I have found this technique limiting due to the small size of the Phast gel and
related small-scale separation of proteins, which tends to result in a messy western blot. I
have had much better success and resolution using minigels and transferring them to
membrane in a separate buffer chamber. However, both techniques are detailed below.
Phast gels
Run a Phast gel as you typically would, following Phast gel protocol. You may
want to run duplicate gels - one to transfer and one to stain in order to compare the
protein gel with your western blot. Use protein standards that are easy to identify on your
gel and on your membrane after transfer (BioRad Kaleidoscope markers are nice as each
protein band is a different color on the gel).
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Monomer Solution
60 g acrylamide
1.6 g bis-acrylamide
water to 200 mL
Tank buffer
30.28 g Tris
155.13 g glycine
10 g SDS
water to 10 L
Water-saturated n-butanol
50 mL n-butanol
5 mL water
mix; use top layer to overlay gels.
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Running Minigels
Assemble minigel using tank buffer (see above) as running buffer in upper and
lower chambers. Typical run for 2 gels is 1 hour at 40 mA constant current. Tracking dye
should run to the bottom of the gel for complete separation. Cooling is optional. Again,
you may want to run duplicate gels, one to stain and one to transfer.
Phast gels
1. Cut PVDF membrane to fit size of actual gel (not stacking gel). Rinse membrane in
methanol briefly, then soak in Towbin transfer buffer for at least 5 min. at room temp.
2. Apply membrane to top of the phast gel in phast gel chamber, taking care to remove
bubbles between membrane and gel. Place the plastic phast gel buffer strip holder
(remove buffer strips, though) over membrane.
3. Transfer simply with heat - set phast system to 60 ºC for 30 min to 1 hr.
4. Soak gel and membrane in methanol and separate.
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The concentration of SDS and methanol can affect transfer. More methanol makes
it more difficult to transfer larger proteins. These concentrations have worked fine for me
with proteins under 100 kDa.
Ponceau S stain
1.0 g Ponceau S
50 mL acetic acid
water to 1 L
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The remaining steps are the same regardless of what gel/transfer method used.
This protocol is a guideline, and many of these recommendations can be optimized for
your particular experiment. The length of blocking time, incubation with antibody,
whether you incubate the membrane with the antibody in the presence of nonspecific
proteins, even the composition of blocking proteins can vary. If you think you have high
background after you visualize your blot, try more stringent blocking and/or washing
conditions. All steps are carried out at room temperature.
TBST
10 mM Tris-HCl pH 8 (5 mL 1M Tris-HCl pH 8)
150 mM NaCl (4.37 g)
0.1% Tween-20 (0.5 mL)
water to 500 mL
TBS
20 mM Tris-HCl (3.2 mL 1M Tris HCl with 0.8 mL 1M Tris base)
150 mM NaCl (1.76 g)
water to 200 mL
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In addition to the links in the text, consult the links below for more information:
http://paris.chem.yale.edu/links.html
The links page of the Schepartz laboratory website.
http://mcb.harvard.edu/BioLinks.html
A good list of biology-related links available through the Department of Cellular and
Molecular Biology at Harvard.
http://www.rcsb.org/pdb/
The Protein Data Bank (PDB) website offers access to the worldwide repository for
processing and dissemination of three-dimensional biological macromolecule structural
data.
http://www.nih.gov/
The National Institutes of Health website contains science news, health resources, and
other scientific resources (U.S. Department of Health and Human Services).
http://www.public.iastate.edu/~pedro/research_tools.html
An extensive list of links to databases, guides, and search and analysis tools of use to the
molecular biologist.
http://www-sci.lib.uci.edu/HSG/GradChemistry.html
A large site with a lot of science information developed by Jim Martindale. There is
chemistry and biochemistry information, periodic tables, and probably more than you’ll
ever have use for.
http://www.webelements.com/
An excellent online periodic table. Clicking on any element will lead you to data about it,
a picture, and related links.
http://chemlab.pc.maricopa.edu/periodic/periodic.html
Another useful periodic table that allows you to choose what properties you want to have
displayed (Phoenix College).
http://www.hhmi.org/research/labsafe/overview.html
This site provides an overview of laboratory safety guidelines, with links to Laboratory
Chemical Safety Summaries (LCSS) and to environmental health and safety departments
of HHMI host institutions.
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MSDS-Search
http://www.msdssearch.com/Default.htm
Chemfinder
http://chemfinder.cambridgesoft.com/
Experimental Calculations
http://paris.chem.yale.edu/extinct.html
A very useful tool on the Schepartz website for calculating the molecular weight of
protein, DNA, or RNA sequences.
http://www.lhup.edu/~rkleinma/Percent.htm
This site provides a clear explanation of how to calculate the percent yield and
underscores the idea of multiple percent yields when dealing with multi-step reactions
(Lock Haven University of Pennsylvania).
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