EXPERIMENTS
IN
CELL BIOLOGY
BIOLOGY 313 LABORATORY MANUAL
WINTER 2002
John E. Rebers, Ph.D.
TABLE OF CONTENTS
Abbreviations, S.I. units of measurement.......................................................................................1
Lab Schedule and Grading..............................................................................................................2
Introduction....................................................................................................................................3
Breakage Fee..................................................................................................................................3
Lab Supplies...................................................................................................................................3
Safety ...........................................................................................................................................3
Laboratory Reports.........................................................................................................................4
Common Lab Report Errors............................................................................................................8
Late Laboratory Reports.................................................................................................................9
Research Projects..........................................................................................................................10
Graph Paper, Linear .....................................................................................................................17
Graph Paper, Semi-Log................................................................................................................18
LABORATORIES
1.
Introduction to the Light Microscope...............................................................................19
2.
Tissue Culture – Culturing an Insect Cell Line................................................................39
3.
Immunocytochemistry – Staining Nuclear Proteins in Insect Cells.................................43
4.
Tissue Culture – Primary Cultures From Chick Embryos................................................49
5.
Protein and Enzyme Assays Using Cultured Chick Cells.................................................55
6.
Neurophysiology..............................................................................................................61
7.
Spectrophotometric Assay of Mitochondrial Succinate Dehydrogenase..........................73
8.
Cell Motility and the Cytoskeleton..................................................................................89
9.
Actin-Based Cell Motility: Myofibril Contraction...........................................................97
10.
Mitosis and the Cell Cycle.............................................................................................105
Abbreviations, Symbols, SI Units, and Prefixes
Term
Abbreviation or Symbol
alpha (lower case Greek letter)
beta (lower case Greek letter)
gamma (lower case Greek letter)
delta (lower case Greek letter)
delta (upper case Greek letter)
lambda (lower case Greek letter)
meter (length)
centimeter
millimeter
micrometer
nanometer
force due to gravity (used for centrifugal force)
revolutions per minute
gram (mass)
milligram
microgram
liter (volume)
microampere
microliter
milliliter
millivolt
molar (concentration)
millimolar
micromolar
degrees Celsius (temperature)
minute (time)
second (time)
milliampere (current)
millivolt (voltage)
ultraviolet
103
10-2
10-3
10-6
10-9
10-12
10-15
kilo
centi
milli
micro
nano
pico
femto
1
m
cm
mm
µm
nm
g
rpm
g
mg
µg
l
µA
µl
ml
mV
M
mM
µM
ºC
min
sec
mA
mV
UV
k
c
m
µ
n
p
f
LAB SCHEDULE & GRADING
Lab
Date
Topic
Maximum
Points
1
1/18/2002
Introduction to the Light Microscope
15
2
1/25/2002
Tissue Culture – Culturing an Insect Cell Line
5
3
2/1/2002
Immunocytochemistry – Staining Nuclear
Proteins in Insect Cells
5
4
2/8/2002
Tissue Culture – Primary Cultures From Chick
Embryos
5
5
2/15/2002
Protein and Enzyme Assays Using Cultured
Chick Cells
15
6
2/22/2002
Neurophysiology
25
7
3/1/2002
Spectrophotometric Assay of
Mitochondrial Succinate Dehydrogenase
25
3/8/2002
No Class - Mid-semester Break
3/15/2002
Research Project
3/22/2002
Research Project
3/29/2002
Research Project
4/5/2002
Research Project
80
8
4/12/2002
Cell Motility and the Cytoskeleton
5
9
4/19/2002
Actin-Based Cell Motility: Myofibril
Contraction
10
10
4/26/2002
Mitosis and the Cell Cycle
10
Points
acheived
"Overspecialization is to be carefully avoided. The biologist who is interested in cell physiology should
not be a morphologist, or a biochemist: he should not only be capable of using physiological and
biochemical methods as well as the microscope, but he should use them all in attacking his problem.
Neither the variety of methods nor the acquisition of a wide knowledge in very different fields should
frighten him. This is the price to be paid if cell physiology is to progress."
Jean Brachet (1957) Biochemical Cytology, Academic Press, NY
2
INTRODUCTION
Goals of the Laboratory
1)
To help you understand the process of science. The information contained in your text is not
revealed wisdom, but is based upon experimental observations. By conducting your own
experiments, you should gain a better understanding of the experimental basis of cell biology.
2)
To introduce you to the tools and techniques used by cell biologists to explore cell structure and
function. The information presented in the text will be less abstract if you are more familiar with
the techniques used to obtain that information.
3)
To help you learn how to record and present scientific data.
4)
To help you improve your writing skills as you prepare laboratory reports.
Preparation for the Lab
Please read the lab exercise before coming to lab, and review the steps that you will need to take to get
started at the beginning of the lab. Most exercises are cross-referenced to your text, Molecular Biology of
the Cell (MBOC), and you will find it easier to understand the purpose of a particular experiment if you
also read the suggested portion of the text before lab. If you have read the procedure before coming to lab,
you will have an easier time understanding the procedures and can focus on the concepts addressed in that
lab.
Breakage Fee
As noted in the NMU Bulletin (p. 12), there is a breakage fee for any glassware or equipment that costs
more than one dollar. The cost for some items, in particular the microscopes, is quite significant. Other
items, like pipets and burets, are less expensive but still more than you might expect. Exercise caution
when working with the equipment and there should be no problem with the breakage fee.
Lab Supplies
Many labs will require a permanent marking pen that will write on glass or plastic. Sharpie®, Vis-a-Vis®,
or similar pens will work well. Graph paper is required for several lab reports. A sheet of graph paper is
supplied at the beginning of the lab manual that can be photocopied if you choose. Alternatively, if
several students share a pack of graph paper the price will be about the same.
Safety
Several laboratories require the use of toxic compounds. Because of this, no eating or drinking is
permitted in the lab. If you bring an empty pop can to lab with you, please store it on the floor so I won't
think that you are drinking from it. Please store coats and books not required for the laboratory on the
racks at the back of the room. Specific safety hazards (for example electric shock or strong acids) will be
described in the description for that lab. Quizzes or exams may ask you to explain why a particular
reagent or procedure presented a hazard, and what could be done to avoid that hazard.
3
WHY ARE THERE LAB REPORTS?
The purpose of lab reports is to help you learn how to:
1) Organize and interpret your data.
2) Present the data clearly, using graphs and tables as appropriate, and
3) Write clearly and concisely, using specific information to support conclusions.
Only a few of the students who take BI 313 will go on to pursue a career as research scientists, where
writing lab reports and research papers is part of the job. However, whatever vocation you pursue, writing
clearly and concisely and making logically supported arguments will be useful to you. Writing did not
end when you completed EN 211 — it will be with you for the rest of your life. The lab reports in BI 313
provide an opportunity to practice and improve your writing skills.
GUIDELINES FOR LAB REPORTS
These guidelines are intended to provide some general help in writing up lab reports for BI 313. Specific
items to be covered in the reports, and occasional exceptions from the format described below, will be
given in lab.
STYLE
Since you are describing the results of experiments you completed in lab, you should use past
tense. Present tense may be used for general statements - however, your report should be specific rather
than general whenever appropriate. Strive to make your report clear, concise, and complete. A useful
reference for hints about writing style is The Elements of Style, by W. Strunk & E.B. White (catalogue #
PE1408, S772; shelved in the reference section). This book describes the most common writing errors and
how to avoid them.
FORMAT
Unless otherwise instructed, the reports should be written in the style used for a scientific paper,
including the following sections: title, introduction, methods, results, discussion, and references. Each
section except the title should be preceded by a heading that identifies the section (for example:
Introduction). The report should be written in a narrative form, with answers to discussion questions
given in a paragraph format, making it clear what issues are being addressed in each paragraph. Do not
simply provide answers to questions from the lab manual that cannot be interpreted out of context, and do
not repeat questions verbatim from the lab manual. You may have used different report styles in the past.
However, just as the editor for a scientific journal will require authors to follow a particular format, I will
require students to follow the format outlined above. Specific exceptions (for example, sections not
required) will be noted in lab. Points will be deducted for not following the specified lab report format. If
reports are printed, please double-space; if reports are handwritten they must be clear and legible.
4
GENERAL GUIDELINES
1) The report should be written in your own words, without using direct quotations from the lab manual
or textbook. If specific, detailed information is included, it is appropriate to quote briefly, but citations
giving the reference quoted should be used.
2) Since you are describing what you did in the lab, the report should be written using past tense.
3) Be specific, rather than general, as much as possible. Your report should focus on describing the
experiments you carried out. It may be useful to refer to other experiments or general information to
provide necessary background for your report, but the principal emphasis should be a description of your
experiments.
TITLE
The title should be short and specific, giving the reader a clear idea of what was accomplished in the
laboratory.
INTRODUCTION
The introduction to the lab report should answer two important questions. First, what did you do? Second,
what is the general significance of the results? Provide a brief description of what was done in the lab, and
what tissues or cells were used, and a short summary of the results. It is also appropriate to provide a
summary of results from previous research done by others and explain how your experiments relate to
these previous results. Remember that references should be provided when discussing the work of others.
The length of the introduction to scientific papers varies depending upon the journal publishing the paper;
your introduction should be 1-3 paragraphs long. The maximum length for the introduction permitted is
one typewritten page, double-spaced. Note that some of the information requested here is included in an
"abstract" or "summary" section in some journals. However, for your report, the introduction should
provide this information.
METHODS
If you used previously published methods to perform the experiments, it is not necessary to provide
complete details for the experiments. Instead, one might report "Mitochondria were isolated from rat liver
using the differential centrifugation protocol described by Smith (1990)"1. However, your methods
section should provide enough information so that a reader familiar with the techniques used will
understand how the experiment was performed without referring back to the lab manual or published
experiments. Remember that one of the key purposes of the methods section is to provide enough
information so that someone else could repeat your results. Therefore, it is especially important to note
any details required to reproduce the experiment under the same conditions you used. It is also important
that the methods section provide a clear description of the overall experimental design. The methods
section should not be a complete recapitulation of the procedures listed in the lab manual, nor should it be
a single sentence saying "See lab manual for methods." A methods section will not be required for all
reports; you will be informed in lab when a methods section is not required.
RESULTS
1
This is an example of the proper way to cite references in the text, but this
is not an actual reference. Be sure to use an appropriate reference and include
that reference in the bibliography when preparing your report.
5
Always report exactly what you observed in the lab. Sometimes the data collected will seem a bit strange,
or differ dramatically from the expected values given in the lab manual or text. You should still report
your own observations, but point out any obvious discrepancies in the discussion. If you can think of a
possible reason for the discrepancy, point this out as well. Remember that the units of measurement
should always be clearly indicated for any data presented.
The results section must include a narrative to provide context for any tables or graphs. Although it
is often most appropriate to include some results as tables or graphs, you also need to include a few
sentences explaining what experiment was done to obtain the data, and how that data fits into a broader
perspective. The narrative portion of the results should clearly explain your experimental objectives –
why did you collect the data? The intent here is not to discuss the overall significance, which is done in
the discussion section, but to provide a context for the results rhat are presented. The narrative portion of
the results should precede any tables or graphs. If you choose, you can integrate tables and figures into the
written narrative of your results section, but you should not begin the results section with a table or graph
without prior explanation of what results are being presented. Alternatively, the tables or figures can be
placed at the end of the report; in this case you should tell where they are [ e.g. see Fig. 4 (p. 9)]. In either
case, be sure to refer to the tables or figures in the narrative of the results and in the discussion. Don't
leave a table or graph as an "orphan" at the end of the report, with no reference to it elsewhere in the
report. You will not receive full credit for your lab report if the results section consists of only
figures and tables, without an associated narrative.
Tables are numerical values presented in columns or rows. Figures are drawings, photographs, or graphs.
Label tables as "Table 1 ..." and graphs, drawings, or photographs as "Figure 1..." Be careful to label all
tables and graphs so that it is clear what data is being presented. Tables should have a title, and labels for
the columns and rows of data. Graphs should have a title, and the abscissa and ordinate should be clearly
labeled to indicate the units used and values presented on each axis. All figures and tables should have a
legend that gives a short summary of the data presented in the figure and explains any abbreviations used.
See figures in a scientific journal such as Cell or Journal of Cell Biology for examples of table and figure
legends. Students may elect to use a graphing program, such as Excel, Lotus, or CricketGraph to prepare
graphs to use as figures. If you do so, you are still required to provide an informative title, legends for the
axes, and use reasonable scales and divisions for the axes. Be particularly careful about the latter point,
since the program defaults may give odd settings for the numerical divisions on the axes. An excellent
reference for more information about using graphics programs to present scientific data is: Briscoe, M. H.
(1996). Preparing scientific illustrations: a guide to better posters, presentations, and publications. (New
York: Springer). Values in a table should be arranged in a logical order - usually either ascending or
descending values. It is sometimes more clear to rearrange the values for presentation in a table rather
than showing the values in the order in which they were collected during the experiment.
DISCUSSION
The discussion section provides an explanation of the significance of your experimental results. The
specific data you collected should be related to general principles. Statements made in the discussion
should be supported by referring to specific data presented in the results section. For example, state that
"The results shown in Figure 1 indicate that enzyme activity increased with temperature." If calculations
are needed to draw a conclusion, show the calculations; for repetitive calculations, showing an example is
sufficient. Describe how your results relate to previous results, calling attention to agreement or
discrepancies with expected results. If you suspect a result is in error, provide an explanation in the
discussion section. However, don't focus too much on the negative. First discuss those aspects of your
experiment that worked, and then move on to the unexpected results.
REFERENCES
6
Citations within your report should give the author and date as shown in the examples below. If a
reference has one author, cite as Smith (1973); if there are two authors, cite as Smith and Jones (1985); if
there are three or more authors the citation in the text uses “et al.” (Latin for “and others”), but the listing
in the bibliography should include all authors (see below).
Karpen et al. (1989) described the role of the Drosophila rRNA gene in nucleolus
formation.
A lariat structure was observed during in vitro splicing of human β-globin mRNA
(Ruskin et al., 1984).
A complete bibliography (including titles of papers) should be given at the end of your report. This
bibliography should follow the format used for the citations in the journal Cell, with full titles and all
authors listed. References should be listed in alphabetical order. Some example references are given
below. Note that all authors names are listed (first initials only are used), the full title of the article is
given (in sentence rather than “title case), and journal titles are abbreviated when appropriate.
Example References:
Elgavish, S., Shaanan, B., 1997. Lectin-carbohydrate interactions: different folds, common recognition
principles. Trends Biochem. Sci. 22, 462-467.
Fristrom, J.W., Hill, R.J., Watt, F., 1978. The procuticle of Drosophila: heterogeneity of urea-soluble
proteins. Biochemistry 17, 3917-3924.
Gooday, G. W. (1990). The ecology of chitin degradation. In Advances in Microbial Ecology, (Edited by
Marshall, K.C.) Vol. 11, pp. 387-430. Plenum, N.Y.
7
COMMON LABORATORY REPORT ERRORS TO WATCH FOR
(You will lose points if you make these mistakes.)
1) Use past tense when describing what you did in the laboratory.
2) Scientific names should be italicized or underlined; genus names are always capitalized and
species names never appear without a genus name or an abbreviation for the genus.
3) Proofread your report. If you prepare the report using a word processor, use the spell check
function to check for spelling errors. In any event, you should read over the report to ensure that it
says what you intended. It is acceptable to neatly correct errors after proofreading, rather than
reprinting the report. It is not acceptable to hand in a report that has not been proofread.
4) Data, flagella, nuclei, and mitochondria are all plural nouns. The singular forms of these nouns
are: datum, flagellum, nucleus, and mitochondrion. Be sure to use the appropriate form of the
noun in the context of the rest of the sentence, and be sure that to use the plural form of a verb if a
plural noun is used.
5) Be sure to use the appropriate number of significant figures when recording numerical data and
when showing the results of calculations. Be as accurate as possible in your measurements, but do
not imply accuracy beyond that by reporting too many significant figures. Remember that if
several numbers with differing significant figures are used in a calculation, the result should be
reported with the number of significant figures for the least accurate of the numbers.
6) Numbers less than zero should have a zero preceding the decimal. Write 0.03 rather than .03, for
example.
7) When using scientific notation, numbers should be written as 3.4 x 106. Other formats are
occasionally used by computer programs that cannot display superscripts (for example 3.4e6).
These formats should not be used when preparing a formal report. To make a superscript when
using Microsoft Word, press the control, shift, and plus keys simultaneously; press the same keys
again to toggle off the superscript.
8) The abbreviation for micrometer is µm, not um; for microampere the proper abbreviation is µA,
not uA; and similarly for other values with the prefix “micro”. If your printer cannot print a lower
case Greek µ, the scientific abbreviation for 10-6, use a u, but add a tail to convert the u to a µ
afterwards.If you are using Microsoft Word, Greek characters are inserted by using InsertSymbol. Make sure that the box on the left (Font) reads normal text, then scroll down to “Basic
Greek” using the box on the right labeled “subset”. You can use the same procedure to insert
special characters such as a degree sign.
9) The results section of your report must include a narrative, and not simply be a collection of
tables and figures.
8
GRADING
Point values for each laboratory report are given on page . Specific topics to be covered in the lab report
are mentioned at the end of the lab exercise. The reports should be consistent with the general guidelines
given above. In addition, the reports should use complete sentences and otherwise be grammatically
correct. If multiple errors in grammar or spelling occur in the report, a 10% deduction will be taken from
the grade. Failure to use past tense appropriately will also result in a 10% deduction from the grade.
PLAGIARISM
Laboratory reports and other course assignments should be your own work. If any portion of a report or
other assignment is copied from another source without providing proper attribution, no credit will
be given for that report. Data from collaborative experiments may be shared, of course, but you should
always indicate if another individual collected the data. If two students hand in identical reports, neither
student will receive credit.
PENALTIES FOR LATE LAB REPORTS
1-7 days late:
10% deduction from grade
8-14 days late:
20% deduction from grade
15-21 days late:
40% deduction from grade
22-35 days late: 60% deduction from grade
More than 5 weeks late: not accepted. No lab reports will be accepted after Monday of finals week unless
documentation of a medical or family emergency is provided.
Exceptions may be made to the above for medical or other valid excuses. If you must turn a report in late
and have a valid excuse, please contact the instructor as soon as possible after returning to class. A
revised deadline for submitting the report will then be agreed upon.
If there are any parts of this manual that are not clear, please let me know about these so the problem can
be remedied for the manual next year. Specific comments are much more helpful than general ones. I
would also like to know which lab exercises you found most helpful to understanding material in the
course, or if there were any exercises that you did not find helpful. You will have an opportunity to
comment on the lab during the course evaluation at the end of the year. Since you may find it easier to
make specific comments just after completing a lab, a critique sheet is provided at the end of each
laboratory. These sheets can be returned in lab, or given to the Biology secretary (NSF 2001)) if you
would prefer to remain anonymous.
9
Research Projects
Part of the cell biology laboratory will consist of a research project experience. Four laboratory periods
near the end of the semester will be set aside to work on the projects. Students will work in groups of four
to design and plan experiments, collect data, and present the results of their experiments to the class.
Although the experiments will not begin until the latter half of the semester, you will need to begin
thinking about your experiments during the first half of the semester. This is especially important if you
would like to do a project that will require special chemicals or time to grow plants to use for the project.
This part of the course is intended to help you understand how cell biologists design and test hypotheses
about cell structure and function. At the end of the semester, all groups will present their experimental
results in a joint poster session. What you gain from this portion of the course will depend on what you
put into it. You are not expected to be able to complete the amount of work presented in a typical
scientific paper in only four lab periods. However, you should be able to formulate a simple hypothesis
and collect data to test that hypothesis in the time provided.
The project you design should test a hypothesis in cell biology. Some very reasonable projects would not
be suitable for this course. For example, testing the sensitivity of different strains of bacteria to different
disinfectants or determining the kinds of bacteria found in soil would be projects more suited to a
microbiology course. However, studying chemotaxis or cell motility in bacteria would be appropriate.
Check with your instructor if you are not sure if a particular project would be suitable.
Planning Your Project
The experiments for the research project will be carried out from March 15, 2002 to April 5, 2002.
Students will work in groups of four for the projects (groups of 3 or 5 may be formed, with the permission
of the instructor, if lab section size does not permit division into even groups of four). Students may form
their own groups within a lab section, submitting the names of all students in their group no later than
March 1, 2002. Students who have not chosen a group by this date will be assigned to one by the
instructor. Each group of students must submit a research proposal, which is due on March 20, 2002.
After completing the experiments, students will prepare a poster describing their experimental results.
This poster session will be held on Friday, April 12, 2002.
Students should begin thinking about possible research projects well before the deadlines. In particular, if
you plan to use vertebrates for your research project, the animals to be used and the experimental design
must be approved by your instructor, and an “Application to Use Vertebrate Animals in Research, Testing
or Instruction” must be approved by the NMU Institutional Animal Care and Use Committee. This
approval process typically takes 3 weeks, and may take longer if the Committee has any questions or if
your experimental plan or application need to be revised. This application must first be approved by your
instructor, and then will be sent to the NMU Institutional Animal Care and Use Committee. Although I
will be happy to help with the preparation of this application, it is your responsibility to submit the
application in a timely fashion, no later than February 22, 2002. A copy of the application is available online at <http://www.nmu.edu/www-sam/graduates/guide/form3.htm>.
10
Some suggestions for projects are provided below. Other sources for ideas include:
b
t
1
Look through books in the library for suitable experiments. You can try searching the laboratory
card catalogue under subject for “Cell Physiology”, “Cytological Techniques”, or “Cytology-Laboratory manuals” You can do a title search for “Methods in Enzymology”, a series that
contains many techniques related to cell biology. You can also design your own searches using
the “Guided Search” page, which allows you to combine search terms.
The American Society for Cell Biology has a number of cell biology laboratory exercises on line
- point your Web browser to <http://www.ascb.org/pubs/exercises.html>
Gustavus Adolphus College has a Web page with a Cell Biology manual, at
http://www.gac.edu/cgi-bin/user/~cellab/phpl?index-1.html
Browsing through the lab manual for BI 313 may give you some ideas about how the experiments
performed in class could be extended, using different experimental conditions or different cell
types.
Your lab project should include an original research component, rather than simply repeating an
experiment or lab exercise that has been previously described. The experiments you plan should be
related to cell biology in some fashion. When planning your lab project, be realistic about the amount of
time you have available, and about the equipment and reagents that will be required to complete your
project. Check with your instructor or with the Biology stockroom manager, to see if particular supplies
are available. A limited amount of funding is available to purchase chemicals for your projects. Be sure to
discuss special needs with your instructor well in advance, since it takes 2-3 weeks for materials to be
ordered and delivered. Please note that a list of equipment and reagents must be included in your research
proposal.
Ideas for Experiments
Some general suggestions about experiments that should be feasible in the time allocated for your
experiments are provided below. Note that your project should ask a new question, rather than simply
duplicating an older experiment.
d
In laboratories 1 and 3, the basic procedures for cell culture are introduced, and
in laboratories 2 and 4, the cultured cells are stained with antibodies to show the
location of specific proteins. There are many other experiments that could be
done with the cultured cells. How quickly do the cells divide? What conditions
affect the rate of division? Do the cells divide coordinately or is the time of
division random?
d
We will study mitochondrial succinate dehydrogenase (SDH) activity in lab 7.
Students could design experiments to test the effect of different compounds upon
SDH - for example, what concentrations of a particular compound will cause
competitive inhibition of SDH? Alternatively, the properties of SDH could be
studied in more detail – what is the pH optimum of cauliflower SDH, or how
does the specific activity of SDH from cauliflower compare to that in other
organisms?
11
The enzyme assays you performed in lab were designed to be carried out with a
simple spectrophotometer. The Biology Department has a much more
sophisticated instrument, called a microplate reader, that takes automated
spectrophotometric readings from a large number of samples. There are a variety
of assays that could be performed using this instrument that could be integrated
into a research project. For more information about possible assays, see
[http://www.moleculardevices.com/pages/max_bib1.html].
[
You might find a technique to isolate a specific organelle from a tissue, and then
measure the activity of an enzyme or enzymes found in that organelle. If two
organelles might be found in the same fraction, you could determine the activity
of both enzymes.
o
In many cell types, heat or other stress causes increased synthesis of proteins
called “heat shock proteins”. These proteins can be analyzed using gel
electrophoresis. What temperatures are needed to induce heat shock proteins in a
particular cell type? What are the sizes of these proteins?
p
Many compounds will affect the rate of cell division. Students could incubate
shoots of broad bean (Vicia faba) or other plants in aqueous solutions of different
concentrations of a test compound such as caffeine, and then determine if this has
changed the proportion of root tip cells undergoing mitosis, using the Feulgen
staining technique from lab 10. Since the seedlings will take several weeks to
grow, you must plant the seeds for this experiment in mid-semester. You should
do background research to determine how many chromosomes are present in the
plant you have chosen to study – this will affect how easy it is to observe mitosis
after doing the Feulgen reaction.
a
There are a variety of experiments that could be done with the cell cultures we
set up in the beginning of the semester. What conditions affect the rate of cell
replication? Are there enzymes or proteins that you would expect these cells to
produce under particular conditions? How would you measure the production of
these proteins?
t
We will observe motility in flagellated algae in laboratory 8. It is possible to
remove the flagella and observe their regeneration. A number of experiments
could be done to test the conditions that optimize flagellar regeneration.
Feel free to consult with the instructor for other experimental ideas. Some cell types that are readily
available for experiments are the cultured cells used in the first several lab exercises, cauliflower and
spinach (sources of mitochondria and chloroplasts), yeast cells, flagellated algae such as Chlamydomonas
or Carteria and bacteria such as E. coli. The equipment and reagents used in scheduled labs will in
general be available for your research project. However, you should consult with the instructor to be sure
any equipment besides the compound microscopes used in lab is available. Be sure to check on the
availability of any chemicals needed for your experiments. If a chemical is not already available in the
biology stockroom, you may need to change your experimental plan. Some reagents can be ordered, but
you should plan for 2-3 weeks for ordering and delivery of any special chemicals. A copy of the Sigma
Chemical catalogue is on reserve at the library. You may find it useful to consult this or other catalogues
to determine if a project is feasible or not. Note: Students should not request chemicals or equipment
from the Chemistry Stockroom. Any such requests, if deemed appropriate, must be made by the
instructor.
12
Research Proposal
Each group of students must submit a research proposal, which is due on March 20, 2002. Howeve, if you
would like to use vertebrates in your project, an Application to Use Vertebrate Animals in Research,
Testing or Instruction must be submitted no later than February 22, 2002. A single proposal will be
submitted by each group, and all students in the group will receive the same grade, with a maximum of 25
points awarded. The research proposal should be 3-4 pages long, printed in a 12 pt font. The proposal
should include:
s Title: Brief and descriptive
:
Authors: Names of all students in the group, and their lab section
:
Summary: A one paragraph description of the research you plan to do
:
Background: A concise review of related experiments that have been conducted, to provide
perspective for your experiments
p Research Plan: A description of the experiments you plan to conduct. You should include a
specific list of methods, reagents, and equipment that will be needed. You should have a general
idea of the results expected, and should include a description of the controls that will be used in
your experiments.
y Timetable: Describe what experiments you plan to carry out on weeks 1, 2, and 3.
:
References: Provide citations for all references cited in the proposal. The citations should include
the author(s), date of publication, title, journal where published, volume number, and pages.
Please prepare the bibliography using the format used in the journal Cell. See the “Guidelines for
Lab Reports” earlier in the lab manual for a description of the proper format for citing references
in the text and for formatting references in the bibliography.
i
List of Equipment and Materials Required: This should be on a separate sheet, and should
include a cost estimate for any chemicals not available on campus. Any requests for chemicals to
be ordered must be made far enough in advance for the order to be processed, and should include
a price estimate. There is a catalogue for Sigma Chemical on reserve in the library. You can
check with Ms. Jingfang Niu, Biology stockroom manager, for catalogues from other suppliers.
Poster Presentation
Students will present the results from their projects as a poster, which is a common format for
presenting results at scientific meetings. The poster session will be held on Friday April 12 from 11 AM
until 2 PM. The laboratory will meet as scheduled from 9-11 or 2-4 PM, with a short laboratory session
on cell motility before or after the poster session. After the scheduled laboratory, students will present
their posters in a room to be designated. There are many possible styles of poster presentation. Guidelines
for poster size will be given later in the semester. Keep in mind that the chief goal is to communicate
clearly. Two excellent sources of ideas about preparing posters are Dazzle Em With Style by Robert
Anholt, and Preparing Scientific Illustrations by Mary Helen Briscoe. Both of these books are on reserve
in the library. Your poster will be worth 35 points towards your final grade.
13
EVALUATION
Your grade for the research project will be based upon: 1) the project proposal, which will be
submitted jointly by each group of four students; 2) an instructor evaluation of individual work and group
cooperation; 3) a written abstract describing the poster, which must be completed the day of the poster is
presented (in real life, this must be done several months before the poster...) and 4) a poster presentation
of student results at the end of the semester. Evaluation sheets for each section are given below.
Project proposal...................................................................................................25
Instructor evaluation............................................................................................10
Poster abstract.....................................................................................................10
Poster presentation..............................................................................................35
Total Points.........................................................................................................80
Evaluation - BI 313 Research Proposal
Group # _____ Total Points: _______/25
Names: _______________________________
_______________________________
_______________________________
_______________________________
____
Summary (2 pts)
Clear, concise, and to the point; briefly provides all relevant information.
____
Background (5 pts)
Reviews relevant prior experiments; provides context for proposed experiments.
____
Research Plan (10 pts)
Specific description of experiments to be conducted. Includes projected results and
needed controls. Realistic and innovative. List of equipment and supplies required
included.
____
Time Line (2 pts)
Realistic schedule for experiments to be completed
____
Bibliography (2 pts)
Complete, follows appropriate format.
____
Overall style (4 pts)
Well organized, easy to read. Includes all required information; submitted in a timely
fashion.
Other comments:
14
Poster Abstract Guidelines and Evaluation
The abstract for your poster should be submitted on an 8.5 x 11" sheet of paper, using a 12 point font. The
following information should be provided:
Line 1: Authors and institutional affiliation. Include your group number on this line.
Line 2: Title (In bold face). Title should be concise and to the point, giving a brief, specific, description of
your project.
Line 3: Abstract. A maximum of 400 words. Describe the work completed and how it relates to previous
work. This section serves the same purpose as the introduction to your lab reports. It would be appropriate
to include the abstract on your poster.
Line 4: References. Cite a maximum of three references that provide context for the research reported.
An example of the appropriate format for the poster abstract is provided on the BI 313 web page.
Evaluation Form - Poster Abstract
Group # _____ Total Points: _______/10
Names: _______________________________
_______________________________
_______________________________
_______________________________
_____ Format specified followed (2 pts)
_____ Title clear, concise, and to the point (2 pts)
_____ Abstract clearly describes experiments and their significance. (5 pts)
_____ Relevant references cited. (1 pt)
15
Poster Evaluation - BI 313 Research Project
Group # _____ Total Points ______/35
Names: _______________________________
_______________________________
_______________________________
_______________________________
____
Title clearly and concisely describes research (2 pts)
____
Necessary background provided in a concise fashion (4 pts)
____
Experimental design is sound; hypotheses to be tested are clearly presented (5)
____
Experimental results clearly presented (5)
____
Figures (drawings, tables, or graphs) used appropriately to communicate ideas (4)
____
Poster is easy to read and communicates ideas with a logical flow (5)
____
Overall quality and originality of research presented (5)
____
Overall aesthetics of poster design (5)
Instructor Evaluation Sheet
Name: ____________________________________
Total Points ______/10
Group Number : _________
You will be evaluated based on the following criteria:
 Was a reasonable amount of time spent on the project?
 Did you have a good working relationship with other students on the project?
 Did you do a fair share of work on the project?
 Were laboratory skills demonstrated or developed during the project?
 Were good time management skills used?
Comments:
16
Linear Graph Paper
17
Semi-log graph paper
18
LABORATORY 1
INTRODUCTION TO THE LIGHT MICROSCOPE
Background Reading:
Molecular Biology of the Cell (MBOC), pp 139-155
INTRODUCTION
The microscope is one of the principal tools of a cell biologist. Although a few cells are large enough to
see without a microscope, usually cells are too small to see without magnification. In this laboratory you
will review the principles of operation of the bright-field light microscope, using several different kinds of
samples. You will also learn how to use phase-contrast microscopy, and compare the appearance of
samples viewed with bright-field and phase optics. We will discuss several important factors that affect
microscopic image quality, and view slides and video examples of several types of samples viewed with
different microscopic techniques.
The most obvious factor affecting a microscopic image is the magnification. The lenses in a microscope
make an image larger and easier to see. Modern microscopes are compound microscopes, with
magnification produced by objective lenses near the specimen and ocular lenses near the eye. The final
magnification of the image is the product of the magnification from the two lenses; generally the
magnification of each lens is etched on the side. Two other important factors affecting the microscopic
image are resolution and contrast.
You will probably have examined some or all of the specific cell types used in this exercise in previous
labs. The objectives for this laboratory are to 1) review the use of the microscope; 2) learn how to make
size measurements with the microscope, and 3) learn how to use phase contrast. We will be making size
measurements in several exercises in the future, and will also be working with phase contrast again. Using
specimens you are already familiar with will make it easier to learn these new techniques.
RESOLUTION
The resolution of an optical system defines the closest proximity of two objects that can be seen as two
distinct regions of the image. Another way of thinking of resolution is that it is the amount of detail that
can be seen in a microscopic specimen. Interference effects created by light passing close to the edge of a
solid object can cause two adjacent objects to appear as a single image (see Fig. 4-3 in MBOC). The limit
of resolution of the naked eye is about 0.1 mm, while a high quality light microscope can attain a
resolution of about 0.2 µm. Good resolution means that objects that are very close can be resolved as two
separate images; thus, the smaller the resolution, the better the image. (One potentially confusing point of
jargon is that if a biologist refers to "high resolution" of an image, it generally means that the image has
objects very close together that can be resolved.)
19
The limit of resolution can be calculated using the Abbé equation:
l.r.
0.61
n sin
l.r. : limit of resolution λ : wavelength of light
θ = ½ the angular width of the cone of light collected by the objective lens
In the Abbé equation, the expression (n sin ) is known as the
numerical aperture. The numerical aperture, or N.A., of the
lens is affected by the cone angle of light that can enter the
lens and by the refractive index of the medium the light
must pass through. Figure 1 compares the angular apertures
for an air (dry) and oil-immersion objective. Because of
refraction, the amount of light that can enter the lens is
affected by the refractive index (n) of the medium between
the specimen and the objective lens. If light moves from a
glass slide (n=1.515) to air (n=1.0), the path of the light will
bend. (Remember how the image of a stick appears bent
when the stick is partially submerged in water). With an oil
immersion lens, immersion oil with a refractive index of
1.515 is placed between the specimen and the objective.
This increases the amount of light that can enter the
objective lens, so the working N.A. of the lens is larger.
Figure 1 (Fig. 32 from Microscope
Basics and Beyond, M. Abramowitz
Therefore, the oil immersion lens shown in Fig. 1 will have a
(1985); used by permission.
larger N.A. not only because the angular aperture of the lens
is larger, but also because the refractive index of oil ("n" in
the equation for N.A.) is greater than the refractive index for air.
The highest quality objectives that can be made have an N.A. of about 1.40. If a violet filter is used to
view specimens with relatively long wavelength light ( =400 nm), the resolution will be 174 nm. Lenses
with an N.A. of 1.40 had been made by the nineteenth century. Although there tends to be a direct
correlation between the numerical aperture of a lens and its price, the physical properties of light make it
impossible to get resolution better than 174 nm using visible light, no matter what you are willing to pay
for the lens. Making the image larger will not improve the amount of detail, just as enlarging a
photographic image too much will not increase detail.
Electron microscopes can resolve much more detail than light microscopes because the wavelength of
electrons is much shorter than visible light. In an electron microscope where the electrons are accelerated
in a 100,000 volt field, the wavelength of the electrons is 0.004 nm, and the theoretical resolution would
be 0.002 nm. Practical limitations of lens design and sample preparation limit the resolution of electron
microscopes to about 2 nm. This is about 100 times better than that possible with the best light
microscopes.
Although electron microscopes can resolve much more detail than light microscopes, the light microscope
is still a very important tool in cell biology. Samples viewed with an electron microscope must be
completely dry, since the microscope operates in a vacuum. In addition, the samples must be cut into very
thin sections (50-100 nm), because electrons cannot penetrate thicker samples. The procedures required to
prepare samples for electron microscopy are quite laborious and require that the cells be dead. (A new
instrument, the high voltage electron microscope, can be used to view some living specimens. However,
samples must still be less than 1 µm thick to allow electrons to penetrate.)
20
Light microscopes can be used to view living tissue. Sample preparation for observing fixed and stained
tissue is usually simpler for the light microscope than for the electron microscope. Although the
resolution of light microscopes has not improved since the nineteenth century, several new methods are
available to provide contrast between the sample and the surrounding material.
CONTRAST
If finely ground glass is immersed in water, it will be nearly invisible. Similarly, many living cells are
nearly invisible when viewed with a bright-field light microscope. This is because the sample is the same
color as the surrounding medium; there is no contrast. If a cell with colored organelles, like chloroplasts,
is viewed in a microscope, more detail can be seen because of the color contrast between chloroplast and
cytoplasm. Contrast in a bright-field light microscope is created by differences in the amount of light
absorbed by different regions of the specimen. In the example above, the chlorophyll in the chloroplasts
absorbs most wavelengths of light but transmits green light; therefore, the chloroplasts appear green.
Since most cellular components don't come in convenient colors, a variety of dyes can be used to add
contrast. Although some of these dyes can be used on living cells, most require that the cells be fixed
(killed) before staining. You will have an opportunity to try different techniques for staining specific
cellular components later in the semester.
The phase contrast microscope and the differential interference contrast microscope exploit
interference effects created as light passes through substances of different refractive indices to create
contrast. Other important types of light microscopes are the fluorescence microscope, the dark-field
microscope, and the polarizing microscope. Two specialized microscopic techniques that have been
developed to provide contrast are autoradiography of cellular components, and computer enhanced
video microscopy. You will learn how to use phase contrast microscopy during this lab and use it to
examine different specimens during the semester. Examples of the other types of contrast methods will be
shown in lab and lecture during the semester.
PHASE CONTRAST MICROSCOPY
The most common technique used to provide contrast for unstained objects that do not absorb light is
phase contrast microscopy. This technique produces contrast using special optical components in the
condenser and the lens of the microscope which exploit the interference effects caused by light moving
through an object more slowly than it moves through the adjacent medium.
The phase contrast microscope takes advantage of differences in the refractive index of cellular
components to create contrast by destructive or constructive interference of the light waves that have
passed through the specimen and those waves that did not pass through the specimen. When light moves
from air to a medium like water or cytoplasm, the speed of light is retarded and the wavelength of the
light is reduced. The ratio of the speed of light in air to the ratio of the speed of light in a particular
medium is known as the refractive index, or R.I.
R.I .
Speed of light in vacuum ( "same" as in air )
Speed of light in medium
Since the speed of light in water, oil, or cytoplasm is less than the speed of light in air, all of these fluids
have a refractive greater than one; for instance, the refractive index of immersion oil is 1.515.
21
As light moves through a cell, the waves are retarded; this causes the light that has passed through the cell
to be out of phase relative to the light that has passed on either side of the cell (Figure 2). In Fig. 2, the
Figure 2 (Fig. 29 from Contrast Methods in
Microscopy - Transmitted Light, M. Abramowitz
(1987); used by permission.
Figure 3 (Figure 30 from Contrast Methods in
Microscopy - Transmitted Light M. Abramowitz
(1987); used by permission.
solid line represents an undeviated ray of light that has not passed through the object, and the dotted line
represents a ray of light that passed through an object with a slightly different refractive index and was
retarded by ¼ wavelength. With standard bright-field optics, this difference in wavelength between the
diffracted wave (which passed through) the specimen and the undeviated wave (which passed through the
background) is not sufficient to provide contrast (Figure 3). In Fig. 3, the dotted line represents the
resultant ray from combining the deviated and undeviated rays illustrated in Fig. 2, and the solid line
represents the undeviated rays and background. Note that the amplitude of the rays from the image
(dotted line) and the background (solid line) is nearly identical; therefore, the contrast is low.
The optical components used in phase contrast microscopy result in the diffracted wave and the
undeviated wave being shifted so that they are ½ wavelength out of phase (Figure 4). This causes
destructive interference, as shown by the diminished amplitude of the resultant wave shown in Figure 4 as
compared to the original deviated wave. Thus, the combination of light waves will result in the specimen
image appearing darker.
22
Two components are added to the microscope to provide phase contrast (Figure 5). A phase annulus, in
the condenser, is designed to allow a ring of light (rather than the normal cone of light) to be focused by
the condenser and pass through the specimen.
Figure 4 (Fig. 33 from Contrast Methods in
Microscopy - Transmitted Light M. Abramowitz
(1987); used by permission.
Figure 5 (Fig. 32 from Contrast
Methods in Microscopy Transmitted Light, M. Abramowitz
(1987); used by permission.
The objective lenses used for phase contrast microscopy contain a phase plate at the back focal plane of
the objective. A narrow ring in the phase plate is thinner than the surrounding material. Since undeviated
light which passes through this ring in the phase plate has less material to move through, the undeviated
light will speed up by ¼ wavelength. Light which passes through a specimen is diffracted, and does not
pass through the phase plate. Remember, however, that the diffracted light was retarded by ¼ wavelength.
Therefore, as the undiffracted and diffracted light combine to form an image, they are (¼ + ¼) = ½
wavelength out of phase. This gives rise to the destructive interference noted in Fig. 4 and a dark image
for the specimen.
The amount of contrast generated will depend upon the thickness and the composition of the specimen.
Thick regions of the specimen will retard light more than thin regions. Portions of the cell with high
refractive index (for instance, organelles with a high concentration of proteins or nucleic acids) will retard
light more than portions of the cell with a relatively low refractive index. These differences in cell
composition are difficult to see in bright-field microscopy, which depends on the absorption of light. The
phase contrast microscope takes advantage of destructive interference to create contrast between regions
of differing refractive index.
23
PROCEDURES
Review the diagram of the microscope below. Ask your instructor if you have any difficulty finding a
particular part. There are three different controls that affect the light intensity – the main switch, the light
intensity knob, and the aperture iris diaphragm knob. Please check all of these controls, as well as the
“phase slider” (see below) before concluding that the light bulb is burnt out.
One important part not illustrated on the diagram below is the “phase slider”, which is a rectangular
sliding bar that can be slid into different positions in the condenser. The phase slider is found just below
the stage, and clicks into four different positions. One position, to be used for bright-field microscopy,
allows all the light to reach the specimen. The other three positions have rings (“phase annuli”) that make
hollow cones of light, which are used to help generate phase contrast. Because the phase annulus is
matched in size to the phase plate in each objective lens, the phase slider must be in the proper position to
get good contrast. Your instructor will show you how to make this adjustment.
Figure 6 Olympus CH30 Microscope
24
General rules for working with your microscope.
1.
Use the same microscope for each lab session. If you notice any problem with your
microscope, report it to the instructor at the beginning of the lab. Each student will be
assigned a specific microscope during the first laboratory session.
2.
Use both hands to carry the microscope to your bench, and be careful to avoid bumping
into other students.
3.
Use coarse adjustment only when working with the 10X objective. For higher power
objectives, use only the fine adjustment knob.
4.
Immersion oil should only be used with the 100X objective. Be careful when working
with immersion oil so that the oil does not get on low power objectives or the stage.
Carefully clean the lens (and slide, if appropriate) when you are through with examining
a specimen using immersion oil. If you spilled immersion oil onto the microscope stage,
wipe it off with a Kimwipe or paper towel.
5.
Only use lens paper to clean lenses. Do not use Kimwipes. The coating on the lenses can
be scratched if lens paper is not used for cleaning. Lens paper is available on the front
bench.
6.
Before putting the microscope away, remove slides, return the microscope to the 10X
objective, and return the condenser ring to the "0" position. Return microscopes to the
proper location in the cabinet, with the number on the arm of the microscope facing out.
7.
Return the phase telescope and green filter to the appropriate box on the front bench.
8.
The breakage fee for equipment applies to microscopes. Microscopes cost a lot more than
one dollar.
25
CALIBRATION OF AN OCULAR MICROMETER
The ocular of a microscope can hold an ocular micrometer, a small glass disc etched with a calibrated
scale that is in focus with the object image. Since the ocular micrometer is in the ocular (surprise), the
size of the scale does not change when a different objective is used. Therefore, the linear divisions on the
ocular micrometer are arbitrary, and the ocular micrometer needs to be calibrated. A glass slide with
known distances marked, called a stage micrometer, is used to calibrate the ocular micrometer.
The stage micrometer has a tiny ruler etched onto its surface. We have three different brands of stage
micrometers, and the length of the ruler and the spacing of the divisions is different for each brand. For
instance, the A&O stage micrometer has a 2 mm ruler, with major divisions of 0.1 mm and minor
divisions of 0.01 mm. The B&L stage micrometer has a 1 mm ruler, with divisions 0.01 mm apart on the
left-hand side of the ruler, followed by larger divisions on the right-hand side. Any of the stage
micrometers can be used for the calibration. Check with your instructor if you are not sure of the ruler
length or the division spacing.
The ocular micrometer found in your microscope has 100 divisions. There are numbers on the ocular
scale every 10 divisions. Depending on the specific ocular micrometer you have, the numbers may read
10-100 (so every division corresponds to a number on the scale), or the numbers may read 1-10, with 10
smaller divisions for each numbered division. Therefore, you may read a cell diameter as 25 divisions
while your lab partner observes 2.5 divisions for the same cell. Since the ocular scale is arbitrary, these
differences will not matter - you will simply make measurements of a known distance on a slide to see
how far apart the arbitrary divisions on the ocular are.
The calibration procedure will be done for the 10X, 20X, and 40X objectives. Do not use the stage
micrometer with the 100X (oil immersion) objective. The stage micrometer is thick enough that the 100X
objective could bump into the slide and scratch the lens.
There is an unfortunate duplication of terms here, in that the same word is used for an ocular micrometer
and for micrometer, a unit of distance equal to 10-6 meters. It is usually clear from the context which is
meant, but be aware of the potential for confusion.
Archaic terminology note: In older literature, you may occasionally see the term “micron” used to refer to
microscopic distances. This term was used as shorthand for micrometer, and is no longer in use.
26
CALIBRATION PROCEDURE
1.
Focus on the calibrations of a stage micrometer using the low power (10X) objective, which gives
100X final magnification. Only one ocular has a micrometer; you may need to close one eye to
see the micrometer clearly. (Note that for other applications, you should keep both eyes open
while using a binocular microscope.)
2.
Adjust the position of the stage and rotate the ocular so that the ocular micrometer scale is aligned
with the divisions on the stage micrometer. Adjust the left end of the ocular micrometer scale so
that it aligns exactly with the left end of the ruler etched on the stage micrometer. Make sure the
two rulers are exactly parallel. Using the 100 X final magnification, it should be possible to
measure a 1 mm distance on the stage micrometer with the full set of divisions on the ocular
micrometer. However, it may not require quite the full set of ocular divisions to match up to the 1
mm line - that is, 98 (or 9.8) ocular divisions might be enough. If the ocular divisions do not
extend for the full 1 mm distance on the stage micrometer, it is acceptable to measure a shorter
distance on the stage micrometer, e.g. 0.9 mm.
3.
Determine the length of each ocular micrometer division by dividing the length of the stage scale
(in micrometers) which is covered by the ocular micrometer by the number of small divisions on
the ocular micrometer required to cover this distance. Enter the values on page 31.
4.
Carefully change to the 20X objective and repeat the measurements made in part 3. Since you
increased the magnification, you will no longer be able to cover a full 1 mm distance with the full
ocular micrometer scale. Instead, measure a distance of 0.5 mm (or 0.4 mm) with the ocular scale.
What is the final magnification with this objective?
5.
Change to the 40X objective and repeat the measurements made in part 3 with this objective,
measuring an appropriate distance on the stage micrometer scale. What is the final magnification
of an object viewed with the 40X objective?
6.
Since the working distance for the 100X (oil immersion) lens is too short to fit the stage
micrometer under the objective, calculate the length of each ocular micrometer division by
dividing the value for micrometers per division obtained using the 10X objective by 10. This
should give you a reasonable value for the value for micrometers per division when using the oil
immersion objective. Enter the value on page 31.
27
WET MOUNTS
In a wet mount, a drop of water or stain is used to provide a continuous film between the microscope slide
and cover slip. The sample should be as thin as practical to provide a high quality image. In a thick
specimen, it is impossible to focus on the entire specimen and out-of-focus parts blur the image. It is also
important to avoid air bubbles and folded specimens. The technique for making a wet mount will be
demonstrated at the beginning of the lab.
Onion Epidermis — Bright-field, unstained
1.
Add a small drop of water to the center of a clean microscope slide.
2.
Cut off a section from a fresh onion. On the concave surface of the section, you should see a
tissue-thin layer of cells. Use forceps or a razor blade to peel free this layer and then use a razor
blade to cut out a section about 1 cm square. Use forceps to transfer this piece to a microscope
slide.
3.
Lay the section of tissue down on the drop of water on your slide, being careful to avoid folding.
Add another small drop of water to the top of the onion tissue.
4.
Hold a cover slip at an angle and lay it down on top of the specimen. If you lower the cover slip
carefully, you should avoid air bubbles. If a few form, don't panic — you should be able to see
the sample in spite of them.
5.
Place the slide on the microscope stage and focus using 100X final magnification (10X
objective). You may find it useful to adjust the initial focus on the edge of the cover slip, and then
move the onion tissue to the center of the field of view.
When using bright-field optics, the condenser should be in the "0" position, and there
should not be a green filter on the illuminator.
6.
Change to 200X and 400X magnification. Since the lenses in your microscope are parfocal, the
specimen should remain in focus as you change objectives. Use the fine adjustment knob to adjust
the focus as necessary.
7.
Examine the unstained onion tissue and answer the questions on page 31.
28
Onion Epidermis — Phase-contrast
1. Return to 100X final magnification (10X objective), and find an area where the specimen is flat and
gives a high-quality image. Adjust the focus so this image is as sharp as possible.
2. Obtain a green filter from the box on the front bench and place it on the illuminator.
3. Move the condenser so that it is as close as possible to the microscope stage.
4. Rotate the dial on the condenser to the "10" position.
5. Answer the questions on page 32.
Note: with most phase contrast microscopes, the phase condenser is adjustable, to provide for optimal
alignment between the annulus in the condenser and the phase plate in the condenser (see Fig. 5). The
simple phase contrast condenser in your microscopes cannot be adjusted. If you use a phase contrast
microscope in the future, be sure to ask how to align the phase condenser.
.
Onion Epidermis — Bright-field, stained
1. Place one or two drops of Lugol's iodine solution on a clean microscope slide and add a fresh piece of
onion epidermis. Examine the stained cells with bright-field optics, using 100X, 200X, and 400X
magnification. Remember to remove the green filter and return the condenser ring to "0". You should
adjust the amount of light to get optimum contrast. Answer the questions on page 35.
Onion Epidermis - Drawing and Measurement
1. Draw an onion epidermal cell, labeling the structures that are visible with phase contrast or after
staining, in the space provided on page 34. Your drawing should be labeled with the name of the
organism (in this case, Allium cepa), the type of optics used for observation, the final magnification
(product of the magnification from the ocular and the objective), and any stain used.
2. Measure the diameter of 6 onion epidermal nuclei, and record the values in the space provided on
page 33.
29
PREPARED SLIDES OF BACTERIA
1. Scan a prepared slide of bacteria, first using 100X magnification and then 400X, to locate an area
with a concentration of bacterial cells.
2. Switch to 1000X final magnification (oil-immersion lens, 100X objective) and examine the shape of
the cells.
3. Draw group of cells from the slide in the space provided on page 35. Be sure to label your drawing
with the type of sample or species of bacteria that were on the slide, and with the final magnification
used for the observation.
4. Measure the length of five bacterial cells (all of the same shape) and record the values on page 35.
5. Be sure to clean the oil immersion lens carefully before putting the microscope away.
CLEANUP
1. Wash off slides used for wet mounts and return them to the container marked "used slides".
Coverslips may be discarded in the broken glass box at the front of the room.
2. Return the condenser to the "0" position, put the low-power (10X) objective in place, and return your
microscope to the appropriately numbered space in the cabinet.
3. Leave your bench area clean and neat.
REFERENCES
Abramowitz, M. (1985). Basics and Beyond Series, 1: Microscope Basics and Beyond. (Lake Success,
NY: Olympus Corporation).
Abramowitz, M. (1987). Basics and Beyond Series, 2: Contrast Methods in Microscopy. Transmitted
Light. (Lake Success, NY: Olympus Corporation).
Taylor, D. L., Nederlof, M., Lanni, F., and Waggoner, A. S. (1992). The new vision of light microscopy.
Am. Scient. 80, 322-335.
30
Laboratory 1 Introduction to the Light Microscope
Name _____________________________________
Section _________________
(Friday AM/ Friday PM)
QUESTIONS
CALIBRATION OF OCULAR MICROMETER
Length measured
on stage scale
(mm)
Length measured
on stage scale (µm)
Number of ocular
divisions required
to match distance
on stage
Length,
µm per division
10X objective
(100X final mag.)
20X objective
(200X final mag.)
40X objective
(400X final mag.)
100X objective2
(1000X final mag.)
WET MOUNTS
Onion Epidermis - unstained, bright-field optics
What cell parts were visible in the unstained preparation with bright-field optics?
_________________________________________
Did you need to increase or decrease the intensity of the illumination to observe the nucleus in the
unstained preparation?
_________________________________________________________________________
_________________________________________________________________________
Why?
Laboratory 1
Introduction to the Light Microscope
Section ____________
Name _____________________________________
(Friday AM/ Friday PM)
Remember that you should not use the 100X objective to examine the stage micrometer. Use the procedure
described on page to calculate the appropriate calibration for the 100X objective.
2
31
Onion Epidermis - unstained, phase-contrast optics
What cell parts were visible in the unstained preparation with phase-contrast optics?
_____________________________________________________________________________________
Describe any details visible within the nucleus with phase-contrast optics.
_____________________________________________________________________________________
_____________________________________________________________________________________
Compare the appearance of the unstained onion epidermis with bright-field and phase optics.
_____________________________________________________________________________________
_____________________________________________________________________________________
_____________________________________________________________________________________
_____________________________________________________________________________________
Onion Epidermis - Stained, bright-field optics
What changes did you need to make in the illumination for observation of the stained cells?
_____________________________________________________________________________________
Compare the appearance of the stained onion epidermis observed with bright-field optics to the
appearance of the unstained epidermis using bright-field microscopy.
_____________________________________________________________________________________
_____________________________________________________________________________________
_____________________________________________________________________________________
Compare the appearance of the stained onion epidermis observed with bright-field optics to the
appearance of the unstained epidermis using phase-contrast microscopy.
_____________________________________________________________________________________
_____________________________________________________________________________________
_____________________________________________________________________________________
_____________________________________________________________________________________
_____________________________________________________________________________________
32
Laboratory 1
Introduction to the Light Microscope
Section ____________
Name _____________________________________
(Friday AM/ Friday PM)
Measurement of onion nuclei
Final magnification used for measurements: ________ X
Nuclear diameter (ocular divisions)
______
______
______
______
______
Average: _________
Use the information obtained when you calibrated the ocular micrometer to convert the average nuclear
diameter from arbitrary ocular division units to micrometers. Show your work below.
Average nuclear diameter (µm): ______________
33
Laboratory 1
Introduction to the Light Microscope
Section ____________
Name _____________________________________
(Friday AM/ Friday PM)
Use the space below for a drawing of a typical cell from the onion epidermis. Be sure to label the drawing
as instructed.
34
Laboratory 1
Introduction to the Light Microscope
Section ____________
Name _____________________________________
(Friday AM/ Friday PM)
PREPARED SLIDES OF BACTERIA
Use the space below to draw the bacteria you observed. Label the drawing as instructed.
Measure the length of the longest axis of 5 cells of the same type. Record the result in ocular division
units.
Final magnification used for measurements: ________ X
Measurements (in ocular divisions): _____ _____ ______ ______ _____
Average length, ocular divisions: ________
Average length, micrometers:
________
Use the information obtained when you calibrated the ocular micrometer to convert the average bacterial
cell length from arbitrary units to micrometers. Be sure to use the conversion appropriate for 1000X final
magnification. Show your work below.
35
LAB CRITIQUE SHEET
Laboratory 1 Introduction to the Light Microscope
Clarity of written introduction/background
Length of written introduction/background
Clarity of in-lab introduction/background
Length of in-lab introduction/background
Clarity of procedures
1 2
Unclear
3
4 5
Very
Clear
2
3
4 5
Too long
1 2
Unclear
3
4 5
Very
Clear
2
3
4 5
Too long
1 2
Unclear
3
4 5
Very
Clear
1 2
Low
3
4
5
High
3
4
5
Excellent
1
Too short
1
Too short
Relevance to lecture material
Rate this lab relative to
other labs at NMU
1
Poor
Specific errors in lab manual, or clarifications needed.
Other comments. (Use back if needed)
36
2
Comments
37
This illustration, from Robert Hooke's Micrographia, shows the plans for his lens-grinding
machine and for his setup of the microscope.
Picture and legend from: http://www.princeton.edu/~his291/Microscope.html
38
LABORATORY 2
TISSUE CULTURE – CULTURING AN INSECT CELL LINE
Background Reading:
Molecular Biology of the Cell, pp. 158-162
Many kinds of plant and animal cells and tissues can be grown in culture after isolation from the intact
organism. There are a number of reasons why this might be done. First, when cells are tissues are grown
in culture, it is easier to observe the effects of adding hormones, growth factors, or other compounds,
since the compound in question can be added at a known concentration to the culture medium without
needing to worry as much about metabolic effects or indirect actions mediated by other cells or tissues.
Second, a homogenous population of a single cell type can be prepared, so the characteristics of that
specific cell type can more easily be studied. Third, it is feasible to add DNA to the cells (a process called
transfection) and then study the regulation of genes in the added DNA or the effect of proteins expressed
by those genes. Of course, experiments done with isolated cells or tissues must be interpreted with
caution, since they may not behave in the same way as they would if they were present in an intact
organism.
Several types of tissue culture experiments can be done. Sometimes an explanted tissue is cultured intact.
Early tissue culture experiments used small pieces of nerve cord to observe the outgrowth of axons.
Developmental biologists sometimes culture fragments of embryos to determine the effect of specific
compounds upon early development. More frequently, the tissues are dissociated into single cells. If the
culture is done with cells that were recently dissociated from an intact tissue, the culture is termed a
primary culture. Cells in primary cultures generally divide only a limited number of times, but have
properties closely related to the tissues from which they were dissociated. Alternatively, established cell
lines can be used. In cell lines, genetic changes have occurred so the cells will divide indefinitely. Cell
lines provide a convenient source of uniform cell populations.
One type of cultured cells has been in the news – embryonic stem cells. These are cultured cells
established from human embryos that have the potential to differentiate into a wide range of tissues.
Preliminary results indicate that these cells could be helpful in treating human diseases, including
Parkinson’s disease, congestive heart failure, and diabetes. However, experimentation with these cells is
controversial because of their embryonic origin. For more information, see the BI 313 home page.
In this laboratory, you will work with GV1 cells, an invertebrate cell line established from embryos of the
tobacco hornworm, Manduca sexta. Each group of students will receive a dish of cells that will be
suspended, counted to determine the cell density, and used to establish four new cultures. After allowing
six days for the cells to grow, two of the cultures will be treated with an insect hormone,
20-hydroxyecdysone. This hormone, which regulates molting and metamorphosis in insects, has been
shown to cause growth changes and elongation in GV1 cells. In the laboratory next week, you will
measure the length of control and treated cells, and also stain the cells with antibodies to reveal the
location of specific proteins.
Note: setting up the cell cultures will not take the entire laboratory period. Students are strongly
encouraged to use the time available to establish groups for the research projects later in the
semester and to begin discussing ideas for the projects.
39
Procedure
1. Students will work in pairs for this experiment. It will work best if one student does the
manipulations and the second student reads the directions and assists.
2. Transfer sterile round coverslips into the top row (A) and the third row (C) of a 24-well tissue
culture dish, by using a sterile Pasteur pipet connected to a vacuum line. The cover slips can be
picked up by turning on the vacuum and deposited in the well by turning off the vacuum. Label
the top row 1:5 and the third row 1:10. Label the dish with your initials and date over the bottom
row.
3. Loosen the lid of a bottle of DL complete culture medium.3
4. Transfer 4 ml of DL complete medium into a sterile 15 ml centrifuge tube. Label the tube 1:5.
5. Transfer 4.5 ml of DL complete medium into a second sterile 15 ml centrifuge tube. Label the
tube 1:10.
6. Use a sterile Pasteur pipet and the vacuum aspirator in the hood to remove the old medium from a
flask of GV1 cells.
7. Add 5 ml of fresh DL complete medium to the culture flaks. Pipet up and down to dislodge the
cells. Avoid frothing the medium during this step.
8. Transfer 1 ml of the suspended cells into the tube with 4 ml of DL complete medium, and 0.5 ml
of the suspended cells into the tube with 4.5 ml of DL complete medium. Mix gently.
9. Transfer 0.5 ml into each of the wells in the top row (A) from the 1:5 cell suspension.
10. Transfer 0.5 ml into each of the wells in the third row (C) from the 1:10 cell suspension.
11. Place your culture dish into the plastic container provided. (This will reduce evaporation). Move
the container into the 28ºC incubator. Several groups can share one container.
12.The cells will be grown for 6 days and then treated for 12-24 hours with 10-6 M 20-
hydroxyecdysone (20HE), an insect hormone that regulates molting and metamorphosis. 20HE
causes GV1 cells to elongate and aggregate, and also stimulates the synthesis of selected proteins
in the cells. Cells in odd numbered wells (A1, A3, A5, C1, C3, C5) will be treated with 20E, and
cells in even-numbered wells (A2, A4, A6, C2, C4, C6) will be treated with ethanol as a control.
References
Alberts, B., Bray, D., Lewis, J., Raff, M., Roberts, K., and Watson, J. D. (1994). Molecular Biology of the Cell, 3rd
ed. (New York: Garland)
Lan, Q., Wu, Z., and Riddiford, L. M. (1997). Regulation of the ecdysone receptor, USP, E75 and MHR3 mRNAs by
20-hydroxyecdysone in the GV1 cell line of the tobacco hornworm, Manduca sexta. Insect Mol. Biol. 6, 3-10.
Lynn, D. E., and Oberlander, H. E. (1981). The effect of cytoskeletal disrupting agents on the morphological
response of a cloned Manduca sexta cell line to 20-hydroxyecdysone. Wilhelm Roux's Archives 190, 150-155.
3
DL complete medium is Grace s Insect Cell Culture medium (Gibco/Invitrogen)
modified by the addition of 10% fetal bovine serum, 2% Yeastolate, 2%
lactalbumin hydrolysate, 1% penicillin-streptomycin-neomycin antibiotic
mixture (Gibco/Invitrogen), 200 mg/l L-alanine, 75 mg/l L-leucine, 50 mg/l Lmethionine, 50 mg/l L-tyrosine; pH adjusted to 6.2.
40
LAB CRITIQUE SHEET
Laboratory 2: Tissue Culture – Culturing an Insect Cell Line
Clarity of written introduction/background
Length of written introduction/background
Clarity of in-lab introduction/background
Length of in-lab introduction/background
Clarity of procedures
1 2
Unclear
3
4
2
3
4 5
Too long
1 2
Unclear
3
4 5
Very
Clear
2
3
4 5
Too long
1 2
Unclear
3
4 5
Very
Clear
1 2
Low
3
4
5
High
3
4
5
Excellent
1
Too short
1
Too short
Relevance to lecture material
Rate this lab relative to
other labs at NMU
1
Poor
Specific errors in lab manual, or clarifications needed.
Other comments. (Use back if needed)
41
2
5
Very
Clear
Comments
42
LABORATORY 3
IMMUNOCYTOCHEMISTRY
Staining Nuclear Proteins in Insect Cells
Background Reading:
Molecular Biology of the Cell, pp. 186-188.
In order for cells to be seen under the microscope, there must be contrast between the cells and the
surrounding medium. In order for different regions of cells to be visualized, there must be contrast
between the internal structures and the cytosol. In the first laboratory, you used phase contrast optics to
generate contrast, based upon different indices of refraction. You also observed cells stained with iodine
(the onion cells) or crystal violet (the B. subtilis slides; these bacteria were stained using a procedure
called Gram staining). In these two examples, contrast is created because the colored stain absorbs light
passing through a region and changes the color or reduces the light intensity. There are a wide variety of
stains with only limited specificity that are useful for identifying different cells or tissues. The
hematoxylin stain and the Wright-Giemsa stain used to stain the blood cells are examples of this.
In other cases, the location of a specific group of chemicals can be revealed with a cytochemical reaction.
Procedures are available to stain DNA, RNA, carbohydrates, or lipids. For example, in the last laboratory
you will use a procedure called the Feulgen reaction to stain DNA in root tips from the broad bean, Vicia
faba. The specificity for this reaction is due to a) treatment of the tissue with hot HCl, which produces
reactive free aldehydes from by depurinating the DNA; and b) Schiff’s reagent, which reacts with the free
aldehydes to produce a red-purple product.
Using the appropriate techniques, it is possible to determine the location of specific proteins or nucleic
acids in cells or tissues. Proteins are localized in cells using a technique called immunocytochemistry in
which antibodies that bind to specific proteins are used to stain cells or tissues. Nucleic acids can be
localized using a technique called in situ hybridization (see pp. 307-308 and Fig 7-20, MBOC; we will not
consider this technique further).
In this week’s lab, we will use immunocytochemistry to detect the ecdysone receptor, a protein found in
the nucleus of insect cells responsive to this steroid hormone. This protein is a member of the intracellular
receptor superfamily (see MBOC, pp. 728-731). The cells to be stained are the Manduca sexta GV1 cells
that were cultured last week.
Antigens and Antibodies
An antigen is any molecule that can provoke an immune response. Generally, large, complex molecules
with variable structures make the best antigens. Carbohydrates and proteins can both provoke an immune
response that is highly specific when the immune cells in the body are exposed to a foreign carbohydrate
or protein. Although nucleic acids can provoke an immune response, the response is generally not as
specific, since there is less variety in the three-dimensional structure of nucleic acids than there is in the
structure of carbohydrates and proteins.
Antibodies are proteins produced by B lymphocytes in response to a foreign molecule, or antigen. Part of
an antibody molecule is always the same (the constant region), while part has a variable structure,
allowing it to make highly specific non-convalent interactions with the antigen. (See Fig. 1).
43
A particular macromolecule will have a wide variety of subregions (formally called epitopes) that can be
recognized by antibodies. If a human protein is purified and
injected into mice, the mice will generate antibodies that
recognize different parts of that protein. The serum (liquid
component of the blood) from the mice will contain antibodies
that will bind to different regions of that protein (see Fig. 2), as
well as antibodies that bind to many other foreign molecules (like
proteins found on viruses and bacteria) that the mouse has been
exposed to. All of these antibodies, collectively called antiserum,
can then be purified from the mouse blood and used to detect the
human protein. Provided that appropriate controls are used, it
may not matter that other antibodies are present as well.
If a more specific reagent is desired, monoclonal antibodies that
recognize a specific region of the protein can be generated (see
Fig. 4-65, MBOC). This procedure allows the production of
antibodies that recognize a specific region of a single protein - for
example antibodies that would bind to the protein twist recognized by
antibody 1 in Figure 2.
If a tissue or cell is incubated with a solution of antibodies under the
appropriate conditions, these antibodies will bind very tightly to specific
antigens in the cells. However, there is an additional problem to solve
before the location of the antigens can be visualized. Although
antibodies are relatively large proteins, they are far too small to see with
a light microscope, and even an electron microscope cannot show their
location in the cell, since the antibodies have almost no contrast
compared to the other proteins in the cytosol or cell membranes. An
additional step is needed to produce contrast.
Visualizing Antibodies in a Cell or Tissue
Two common ways to do this are to couple antibodies to a fluorescent tag (immunofluorescent
microscopy), or to an enzyme that will produce a colored product (immunoenzymatic microscopy). It is
possible to label the constant region of the antibody with one of these tags, while leaving the variable
region free to react with the protein of interest. Then the fluorescence or colored reaction product in the
cell or tissue indicates where that protein is located. However, the procedure for labeling the antibody is a
moderate amount of work. Since there are thousands of different kinds of antibodies that can be used to
identify proteins in cells, it would be a lot of trouble to label each one.
This problem can be minimized by using the technique of indirect immunofluorescence. This technique
takes advantage of the fact that the constant region of an antibody has a sequence of amino acids that is
the same in a particular species, but is slightly different in another species. Therefore, if antibodies from
mice are injected into another species, like rabbits, the rabbits will generate antibodies that recognize the
mouse antibodies. Antibodies that recognize specifically the constant region of mouse antibodies that are
already labeled with either a fluorescent or an enzymatic tag can be purchased commercially. This allows
an investigator to use a “sandwich technique”. First, the cell or tissue is incubated with a mouse antibody
that binds to a specific protein. This antibody, called the primary antibody, is not labeled. Excess antibody
is washed off, and the tissue is then incubated with a rabbit antibody that binds to the constant region of
any mouse antibody. This secondary antibody is labeled with a fluorescent or enzymatic tag. Fluorescence
44
microscopy or an enzymatic reaction is then used to reveal the location of the secondary antibody. This
procedure has two advantages. First, a wide variety of labeled secondary antibodies can be purchased at a
moderate price, saving the time and labor of the labeling step. Second, several molecules of secondary
antibody may be able to bind to a single molecule of primary antibody. This provides an amplification of
the signal.
Fixation
Before cells are stained for microscopic observation, the cells are often subjected to a process called
fixation. Fixation:
f Helps cells or tissues adhere to the microscope slide;
H Makes cells more permeable to the stain;
M Cross-links macromolecules within the cell, which helps by:
o Preserving cellular morphology
o Inactivating degradative enzymes (like proteases or nucleases)
o Preventing material from washing out of the cell.
There are several common fixatives, including alcohols (usually methanol or ethanol), acetic acid, and
reactive aldehydes such as formaldehyde and glutaraldehyde. In today’s laboratory we will use
paraformaldehyde (a dry form of formaldehyde) in solution in a buffered saline solution to fix the cells,
and will also treat the cells with Triton X-100, a non-ionic detergent, to make them permeable to the
antibodies.
Finding the optimal fixation and staining procedures is largely empirical, with adjustments needed for
different cells or tissues. One important consideration, both when developing staining techniques and
when interpreting results, is the possibility of artifacts appearing in the specimen. An artifact is a structure
or experimental result obtained due to the method of specimen preparation or the experimental conditions.
Since we are interested in what is found naturally in the specimen, artifacts are at best a distraction and at
worst can mislead the investigator.
REFERENCES
Harlow, E., and Lane, D. (1999). Using antibodies: a laboratory manual, 2nd ed. (Cold Spring Harbor, NY:
Cold Spring Harbor Laboratory Press).
Naish, S. J., Boenisch, T., Farmilo, A. J., and Stead, R. H., eds. (1989). Handbook of Immunochemical
Staining Methods. (Carpinteria, CA: DAKO Corporation).
Roitt, I., Brostoff, J., and Male, D. (1998). Immunology, Fifth ed. (London: Mosby).
45
PROCEDURES
[Adapted from Harlow and Lane (1999); Portable Protocol No. 2, Staining cells growing on coverslips.]
1. Remove the culture medium by aspiration and wash the cells once with phosphate-buffered saline
(PBS). Sterility is not needed for this or later staining steps.
2. Aspirate the wash buffer and fix the cells by adding 4% paraformaldehyde. Approximately 1 ml
should be added to each well – the exact volume is not critical. Incubate in the hood for 10
minutes at room temperature.
3. Remove the paraformaldehyde by aspiration and wash the cells twice with PBS.
4. Aspirate the last wash buffer and permeabilize the cells by adding 0.2% Triton X-100 in PBS.
Incubate for 5 minutes at room temperature.
5. Remove the detergent solution by aspiration. Wash the coverslips in 0.2% Triton X-100 with
three changes over 5 minutes. Drain well but do not allow the specimens to dry. It is important to
proceed quickly and efficiently with the antibody addition in step 6.
6. Add 25 μl goat serum to the coverslips in wells A1, A2, C1, and C2. Make sure the coverslips do
not touch the sides of the wells.
7. Add 25 μl of a 1:5 dilution of the primary antibody to the coverslips in wells A3, A4, C3, and C4.
Make sure the coverslips do not touch the sides of the wells. The primary antibody we will use is
designated 9B9. It is a monoclonal antibody that recognizes the Manduca ecdysone receptor.
8. Add 25 μl of undiluted primary antibody to the coverslips in wells A5, A6, C5, and C6. Incubate
for 60 minutes at room temperature. Note: this 60 minute incubation is an excellent
opportunity to discuss ideas for the research project to be done later in the semester with
other students or your instructor.
9. Wash the coverslips in three changes of 0.2% Triton X-100 in PBS over 5 minutes. After the last
wash, drain well but do not allow the specimens to dry.
10.Apply 25μl of the fluorescently-labeled secondary antibody (conjugated to Texas Red). Incubate
for 20 minutes at room temperature.
11. Wash the coverslips in three changes of 0.2% Triton X-100 in PBS over 5 minutes. Drain well.
12.On a clean and labeled microscope slide, place a drop (approximately 50 μl) of mounting
medium.
13.Remove the coverslip from the dish using fine-tipped forceps and drain the last of the wash buffer
by touching the edge of the coverslip to a clean paper towel. Invert the coverslip and place on the
drop of the mounting medium with the cell side down. Gently lower the coverslip, touching one
edge to the slide next to the drop, then allowing the coverslip to fall on the drop. Allow to air dry
for at least 30 minutes prior to observing.
14. If time permits, we will observe and photograph the slides in this lab period. Otherwise, the slides
will be stored and observed later.
46
LAB CRITIQUE SHEET
Laboratory 3: Immunocytochemistry – Staining Nuclear Proteins in Insect Cells
Clarity of written introduction/background
1 2
Unclear
3
4
5
Very
Clear
Length of written introduction/background
1 2
Too short
3
4
5
Too long
Clarity of in-lab introduction/background
1 2
Unclear
3
4 5
Very
Clear
Length of in-lab introduction/background
1 2
Too short
3
4
Clarity of procedures
1 2
Unclear
3
4 5
Very
Clear
1 2
Low
3
4
5
High
3
4
5
Excellent
Relevance to lecture material
Rate this lab relative to
other labs at NMU
1
Poor
Specific errors in lab manual, or clarifications needed.
Other comments. (Use back if needed)
47
2
5
Too long
Comments
48
LABORATORY 4
TISSUE CULTURE – PRIMARY CULTURES FROM CHICK EMBRYOS
In the second laboratory, you worked with a cell line that had been established by homogenizing embryos
of the tobacco hornworm, Manduca sexta, followed by selecting cells that were able to continue dividing
in culture. There are thousands of different cell lines that have been established, both from vertebrate and
invertebrates, as well as from a variety of plants. To get some idea of the different kinds of cell lines
available, visit the American Type Culture Collection web page at http://www.atcc.org/ and choose “Cell
Biology” in the “Search a Collection” query box.
All of these cell lines started out as a primary cell culture. Tissues are dispersed into a single cell
suspension (or in some cases cultured as aggregated tissues) in a medium that contains the nutrients and
growth factors needed to support growth and division. Not all cells in a tissue are capable of growing in
culture. In addition, many culture procedures select for those cells that can attach to the surface of the
culture vessels used. Thus, culturing selects those cells that can tolerate the specific culture conditions
used.
More significantly, most cells in culture will divide a limited number of times, and then will stop
dividing. This is not a matter of nutrients running out, since the same cells could divide when fewer
generations in culture had elapsed. The number of divisions varies from one cell type to another, but
approximately 50 divisions is fairly typical. Cell biologists are currently investigating the factors that
limit the total number of cell divisions, and are also trying to understand how cells “count” the number of
divisions that have elapsed.
Although established cell lines are often more convenient to work with, there are a number of reasons
why a cell biologist might choose to work with a primary cell culture instead. One reason is that many
(and perhaps all) cell lines that are capable of indefinite growth in culture have undergone genetic
changes to allow this indefinite growth. For at least some cell lines that have been studied, these changes
are very similar to those changes that take place when a tumor develops. Although this presents
fascinating opportunities for investigating the nature of cancer, it also means that cell lines have some
drawbacks if the primary goal is to understand normal cell behavior. Those scientists working with
established cell lines are aware of this issue and interpret their results with caution. However, for some
studies, a primary cell culture that has not undergone the genetic changes required for continuous growth
may be preferred.
Another reason a scientist may choose to work with a primary cell culture is that there may not be an
established cell line from the species or tissue of interest, or those cell lines that are available may not
exhibit the behavior or characteristics of interest for a particular study. In this case, it may be preferable to
establish a primary culture.
In today’s laboratory, two different primary cell cultures will be established. Half of the class will
establish fibroblast cultures from minced chick embryos. The other half of the class will establish heart
cell cultures. Be careful to use aseptic technique to avoid contaminating your culture. We will allow the
cultured cells to grow for 1 week, and then will stain the cells to reveal the presence of tubulin, a
cytoskeletal protein.
49
Procedure – Establishment of a Primary Culture of Chick Fibroblasts
The protocol below is from the Cell Biology Laboratory Manual by Dr. William Heidcamp at Gustavus
Adolphus College [http://www.gac.edu/cgi-bin/user/~cellab/phpl?index-1.html]
Procedures should be carried out in a laminar flow hood to minimize the risk of contamination. Be sure to
use proper aseptic technique.
1. Candle an 8 day old egg to ensure that it is alive. This is easily accomplished by holding the egg
in front of a bright light source; the embryo can be seen as a shadow. Circle the embryo with a
pencil.
2. Place an 8-10 day chicken egg in a beaker with the blunt end up, and wash the top with a mild
detergent, followed by swabbing with ethanol.
3. Carefully puncture the top of the egg with the point of a pair of sterile scissors and cut away a
circle of shell, thus exposing the underlying membrane (the chorioallantois).
4. With a second pair of sterile scissors, carefully cut away and remove the chorioallantoic
membrane, exposing the embryo.
5. Identify and carefully remove the embryo by the neck, using a sterile metal hook or a bent glass
rod, and place the embryo in a 100 mm Petri dish containing phosphate buffered saline (PBS).
Wash several times with PBS by transferring the embryo to fresh Petri plates. After removal of all
yolk and/or blood, move the embryo to a clean dish with PBS.
6. Using two sterile forceps, remove the head, limbs, and viscera. Be sure to remove the entire limb
by pulling at the proximal end. Move the remaining tissues of the embryo to yet another dish and
wash with PBS.
7. Mince the embryo finely with scissors and transfer the minced tissue to a flask containing PBS.
Allow the tissue pieces to settle.
8. Remove the PBS with a sterile pipette and add 25 ml of trypsin, a proteolytic enzyme. Stir the
solution gently at 37° C for 15-20 minutes.
9. Allow the larger, undigested tissue pieces to settle and decant the supernatant into an equal
volume of Minimal Essential Medium (MEM) + 10% Fetal Bovine Serum (FBS). FBS contains
protease inhibitors which will inactivate the trypsin.
10.Centrifuge the cells in MEM at 1000 rpm for 10 minutes in a standard clinical centrifuge.
Remove the supernatant and resuspend the pellet in 25 ml of fresh MEM + 10% FBS.
11.Remove 0.1 ml of the culture and determine cell concentration and viability using Trypan blue
staining. A handout will be available explaining this procedure.
12.Seed six 25 cm2 plastic culture flasks containing 25 ml of MEM + 10% FBS to a final
concentration of 105 cells/ml.
50
13. Label and place your cultures in the tissue culture incubator at 37° C and examine daily for cell
density and morphology.
14.Note any changes in the color of the media. The tissue culture media used for the chick cells has a
pH indicator (Phenol Red) added in order to check on the growth of cells. The media initially is a
cherry red (with slight blue haze) and turns orange and then yellow as the cells grow, thereby
reducing the media. Should this color change occur within 24 hours, the culture is most likely
contaminated and should be disposed of.
15. Examine the cultures using an inverted phase contrast microscope. This will allow observation of
the cells without opening or disturbing the growth.
Procedure – Establishment of a Primary Culture of Chick Heart Cells
The protocol below is adapted from Laboratory Exercises in Cell Biology (1989) I.R. Schmoyer and J.R.
Vaughan, Hunter Textbooks, Winston-Salem, N.C.
Preparation before dissection of the chick embryo
P
Place a sterile 120 mm Petri dish under a dissecting scope
P
Fill four 35 mm Petri dishes half-full with warm phosphate-buffered saline (PBS)
F
Fill an Erlenmyer trypsinizing flask with 3 ml of trypsin-EDTA solution
F
Place the pipets and dissecting instruments in an accessible location
P
Add 4.5 ml modified Eagle’s medium (MEM) in each of six 25 mm2 tissue culture flasks
The dissecting microscope, the 120 mm Petri dish, one of the 35 mm Petri dishes, and the dissecting
microscope can be out on the open bench. The remaining reagents should be in the culture hood.
Steps 1-4 can be carried out on the open bench, working carefully to avoid contamination.
Steps 5-16 should be carried out in a laminar flow hood
1. Candle an 8 day old egg to ensure that it is alive. This is easily accomplished by holding the egg
in front of a bright light source; the embryo can be seen as a shadow. Circle the embryo with a
pencil.
2. Wash the egg with a mild detergent, followed by swabbing with 70% ethanol.
3. Crack the egg against a clean, blunt surface and pour it into the top half of a 120 mm sterile Petri
dish. Discard the shell. Moisten the embryo with warm PBS as needed to prevent drying.
4. Remove the embryo and place it in the bottom half of the 120 mm sterile Petri dish. Using the
dissecting microscope, quickly remove the heart by pulling open the chest cavity with two sterile
forceps and place it in the first of the series of 35 mm Petri dishes.
51
5. Transfer the 35 mm dish with the heart to the laminar flow hood. Squeeze the heart to remove any
trapped blood and carefully remove any attached vessels. Transfer the heart to the second and
then the third 35 mm dish, shaking gently to remove blood and any other loose material.
6. In the third dish, the heart should be cut into 1 mm cubes with a sterile scalpel and forcep. Try to
avoid tearing the tissue. Gently swirl the dish.
7. The washed heart fragments from several groups can be pooled and added under sterile
conditions with a forcep to 3 ml of prewarmed trypsin solution being stirred in a 50 ml Erlenmyer
flask. Stir the fragments until the solution begins to appear cloudy, but no more than 10 minutes.
8. Add 3 ml of warmed fetal bovine serum to stop the activity of the trypsin.
9. Allow the large fragments to settle for a minute or two, and then withdraw the dispersed cells
using a sterile 5 ml pipet. Transfer the cells to a sterile 15 ml centrifuge tube.
10. Centrifuge for 6 minutes at 200 g at room temperature.
11. Remove the supernatant and suspend the pellet in 5 ml MEM.
12. Centrifuge for 6 minutes at 200 g at room temperature.
13. Remove the supernatant and suspend the pellet in 3 ml MEM.
14. Centrifuge for 6 minutes at 200 g at room temperature.
15. Remove the supernatant and suspend the pellet in 3 ml MEM. Add additional MEM until the
suspension just barely appears turbid.
16. Transfer 0.5 ml of the cell suspension to the culture flasks containing 4.5 ml of MEM.
17.Incubate the cells overnight at 37ºC in a CO2 incubator set to provide 5% CO2.
18. Observe the cells after 24 hours of growth. Many cells will have attached and begun to divide and
grow out from the point of attachment. Remove the old medium and replace with 5 ml of fresh
MEM.
52
LAB CRITIQUE SHEET
Laboratory 4: Tissue Culture – Primary Cultures from Chick Embryos
Clarity of written introduction/background
1 2
Unclear
3
4
5
Very
Clear
Length of written introduction/background
1 2
Too short
3
4
5
Too long
Clarity of in-lab introduction/background
1 2
Unclear
3
4 5
Very
Clear
Length of in-lab introduction/background
1 2
Too short
3
4
Clarity of procedures
1 2
Unclear
3
4 5
Very
Clear
1 2
Low
3
4
5
High
3
4
5
Excellent
Relevance to lecture material
Rate this lab relative to
other labs at NMU
1
Poor
Specific errors in lab manual, or clarifications needed.
Other comments. (Use back if needed)
53
2
5
Too long
Comments
54
LABORATORY 5
PROTEIN AND ENZYME ASSAYS USING CULTURED CHICK CELLS
Introduction
In last week’s laboratory, primary cultures of chicken fibroblasts and heart cells were established. This
week we will harvest cells from the two different cultures, prepare cell extracts, and determine the amount
of the enzyme acetylcholinesterase in the extracts. Acetylcholinesterse is expressed in muscle tissue, so
one would expect the enzyme level to be higher in the heart cell cultures. To control for differences in cell
size or number in the cultures used to make the extracts, we will also determine the total protein
concentration.
Preparation of Cell Extracts
Follow the procedure described below using either a flask of cells with a fibroblast culture or a flask of
cells with a heart cell culture. Since the cells are to be lysed, you do not need to use aseptic technique.
This procedure is adapted from Gasque (1989).
1. Uncap the flask of cells and decant the medium into a waste container with disinfectant. Add 5 ml
of pretrypsinizing solution (0.8% NaCl; 0.03% KCl; 0.138% Na2HPO4; 0.2% KH2PO4; 0.05%
Na2EDTA) to the flask and recap the flask.
2. Hold the flask flat so the solution flows gently over the monolayer of cells, Rock gently from side
to side for 10-15 seconds to rinse the cells. This step removes residual serum, which would inhibit
the trypsin; the EDTA chelates Mg2+ and Ca2+, which are important in the adhesion of cells to one
another and the plastic substrate.
3. Uncap the flask and decant the pretrypsinizing solution into a waste container.
4. Repeat steps 2 and 3.
5. Transfer 1 ml of trypsin-EDTA solution (0.05% trypsin, 0.02% Na2EDTA in PBS) into the
culture flask. Rock the flask gently from side to side for 15-30 seconds to expose the cells to the
enzyme. Uncap the flask and drain excess trypsin into a waste container, leaving behind a thin
film of solution. Recap the flask.
6. Incubate the flask at 37ºC for 5-15 minutes, until the cells round up and begin to detach from the
substrate.
7. Rap the flask firmly against the palm of your hand to assist in detaching cells and to disrupt cell
clumps.
8. Transfer 3 ml of PBS into the flask. Aspirate the medium up and down with a 5 ml pipet, rinsing
off the bottom of the flask.
9. Transfer the cells into a 15 ml centrifuge tube. Centrifuge at high speed in a clinical centrifuge for
5 minutes.
10. Pour off the supernatant and add 3 ml PBS. Resuspend the cells and centrifuge at high speed in a
clinical centrifuge for 5 minutes.
11. Suspend the cells in 1 ml of PBS containing 0.25 mM EDTA and 0.5% Triton X-100.
55
12. Transfer the suspended cells to a glass homogenizer. Homogenize the cell with 5-10 strokes of
the pestle.
13. Use a Pasteur pipet to transfer the homogenate to a 1.5 ml microcentrifuge tube.
14. Centrifuge for 5 minutes at 4ºC at full speed in a microcentrifuge.
15. Store the homogenate on ice until ready for the protein and enzyme assays.
Measurement of Total Protein Concentration
We will be using a commercial protein assay to determine total protein concentration in the cell
extracts. This assay, sold by Bio-Rad, is based upon the method of Bradford (1976). A solution of
Coomassie Brilliant Blue G-250, which is light brown in color, is added to a test protein sample.
When the dye binds to protein, it shifts to a blue color, with an absorption maximum at 595 nm. The
intensity of the blue color is proportional to protein concentration. Standards of known protein
concentration are included to provide a relative measurement of protein concentration.
Before beginning the assay, take 10 μl of each unknown protein sample to be measured and add it to
40 μl of PBS. This will provide a 1:5 diluted protein sample to use in the assay in case the original
sample is outside the linear range of the assay.
Protein Assay Procedure
1.
2.
3.
4.
Add 10 μl of PBS to wells A1, A2, A3, and A4 of a microtiter plate. These samples
will be used as a blank for the spectrophotometer reading and as a 0 mg/ml protein
standard.
Transfer 10 μl of protein standards with concentrations ranging from 0.2 to 1.0 mg/ml
into wells A5-A12 of the microtiter plate. Each standard should be loaded in duplicate.
Exact concentrations for the standards will be provided in class. Record these values in
the grid below.
Complete the grid on page 58 with the cell extracts to be assayed. Each extract should
be assayed in duplicate. You should assay the undiluted samples as well as the 1:5
diluted samples. Transfer 10 μl of the cell extracts to be assayed into wells after
completing the grid. There should be enough space in the microtiter well for all
samples from the class.
Add 200 μl of diluted Bio-Rad protein assay reagent into each well containing samples
or standards. Empty wells do not need reagent.
5. Incubate at room temperature for at least 5 minutes but no longer than 1 hour. While
the samples are incubating, set up the microplate reader to measure the absorbance at
595 nm.
56
Acetylcholinesterase Assay
1. Add 290 μl of assay buffer [0.01% DTNB, 5,5’dithiobis(2-nitrobenzoic acid) in 0.05 M
phosphate buffer, pH 7.4] to duplicate wells for each extract to be assayed. Include two wells for
sample blanks and two wells for positive controls. Complete the grid on p. 58 to indicate what
samples are to be placed in each well. Undiluted samples and 1:5 diluted samples should both be
assayed. There should be enough wells in the microtiter plate to assay all the class samples in one
run.
2. Add 6 μl of cell extract to each well. The sample blanks will receive PBS without any cell
extract.
3. Add 6 μl of acetylthiocholine iodide substrate solution (2.5 x 10-2M acetylthocholine iodide in
PBS). It is important to add this solution to all wells quickly and begin reading immediately
afterwards.
4. Measure the absorbance at 405 nm at 1 minute intervals for 20 minutes. Be sure the microplate
reader is programmed to begin these readings before adding the substrate.
Laboratory Report
Answer the questions below. A formal report is not required for this laboratory.
1. Calculate the specific activity for each extract. The specific activity is ΔA405 min-1 (mg protein)-1.
Instructions on how to calculate the specific activity will be distributed in class.
2. Is there a difference in acetylcholinesterase activity between the fibroblast cultures and heart cell
cultures? If so, how much?
3. What is the function of acetylcholinesterase? Why is this enzyme expected to be found in muscle
cells and not in fibroblast cells?
4. Why was it important to determine the total protein concentration in the extracts, rather than
simply comparing the acetylcholinesterase activities?
References
Ashour, M. B., Gee, S. J., and Hammock, B. D. (1987). Use of a 96-well microplate reader for measuring
routine enzyme activities. Anal. Biochem. 166, 353-360.
Bradford, M. M. (1976). A rapid and sensitive method for the quantitation of microgram quantities of
protein utilizing the principle of protein-dye binding. Anal. Biochem. 72, 248-254.
Gasque, C. E. (1989). A Manual of Laboratory Experiences in Cell Biology, (Dubuque, IA: Wm C Brown.
57
Protein Assay Template
1
2
3
4
5
6
7
8
9
10
11
12
9
10
11
12
A
B
C
D
E
F
G
H
Acetylcholinesterase Assay Template
1
2
3
4
5
6
A
B
C
D
E
F
G
H
58
7
8
LAB CRITIQUE SHEET
Laboratory 5: Protein and Enzyme Assays Using Cultured Chick Cells
Clarity of written introduction/background
1 2
Unclear
3
4
5
Very
Clear
Length of written introduction/background
1 2
Too short
3
4
5
Too long
Clarity of in-lab introduction/background
1 2
Unclear
3
4
5
Very
Clear
Length of in-lab introduction/background
1 2
Too short
3
4
5
Too long
Clarity of procedures
1 2
Unclear
3
4
5
Very
Clear
1 2
Low
3
4
5
High
3
4
5
Excellent
Relevance to lecture material
Rate this lab relative to
other labs at NMU
1
Poor
Specific errors in lab manual, or clarifications needed.
Other comments. (Use back if needed)
59
2
Comments
60
LABORATORY 6
NEUROPHYSIOLOGY
Students will need to bring a laptop computer to class for this laboratory. A limited number of
computers will be available in the laboratory for students to use. The experiment can be done by
students working in pairs – if you do not have a computer you can bring to class, make
arrangements to work with another student or to share the cost of renting a computer for the day
of class with another student.
Background reading:
Required:
Chapter 11, Molecular Biology of the Cell, 523-535
Membrane potential and ion channels
Voltage-gated ion channels and the action potential
Optional:
Stevens, C.F. (1979). The neuron. Sci. Am. 241, 54-65.
Keynes, R.D. (1958). The nerve impulse and the squid. Sci. Am. 199, 83-90.
The information below provides background and explains how to use the computer program, Neurosim
for Windows, which is designed to simulate a broad range of experiments in neurophysiology. The
specific experiments you will be required to complete are described below. You will need to carefully
read the lab and assigned reading before coming to lab in order to complete the experiments in three
hours. If you choose, you can complete additional experiments in neurophysiology for extra credit (see p.
69)
BACKGROUND
Although a voltage difference, or membrane potential, can be measured across the plasma membrane of
all living cells, some cells possess the special property of being electrically excitable. When the plasma
membrane is depolarized, voltage-gated ion channels in the membrane can respond by opening and
allowing a further depolarization of the membrane. This depolarization spreads as a wave of excitation
called the action potential. Both muscle cells and nerve cells have the special ion channels that allow them
to be electrically excitable; all cells possess the Na+/K+ ATPase and K+ leak channels that are used to
establish the membrane potential.
During the 1940s and 1950s, work with the giant axon of the squid Loligo provided great insight into the
way in which action potentials are conducted. The "jet propulsion" which squids use as an escape
mechanism is controlled by two sets of giant nerve fibers (see diagrams in Keynes, 1958). The axons of
these fibers are up to a millimeter in diameter and several centimeters long — pretty impressive for a
single cell. This large size allowed recording electrodes to be inserted into the cell so that changes in
membrane potential could be measured. Besides its large size, the squid giant axon has the virtue of being
sufficiently robust to withstand extrusion of the normal cytoplasmic contents so that the ion
concentrations inside and outside the neuron could be changed experimentally. Panel 11-3 (p. 531,
61
MBOC) summarizes some experiments done with the squid giant axon. A.L. Hodgkin and A.F. Huxley
were awarded a Nobel prize for this work in 1963.
Since squid are not found in Lake Superior, we will be using a computer model to simulate the behavior
of the squid giant axon. Neurophysiologists generally use several different pieces of equipment for their
experiments: a microscope and micromanipulator, to observe the neurons and place recording and
stimulating electrodes in appropriate places within the neuron; a stimulator, which delivers stimuli of
varying strength and duration to the neuron, and an oscilloscope, which is used to record very rapid
changes in voltage or current across the plasma membrane. The program you will be using in this lab,
Neurosim for Windows, will allow you to observe the electrical changes which would occur with an
isolated giant axon from the squid Loligo.
USING THE NEUROSIM FOR WINDOWS PROGRAM
We will be using a computer program available on the campus network for this laboratory. This program
can used anytime you are on campus and logged onto the network, but is not accessible by dial-in access.
Since nearly all computer labs on campus have been eliminated, we will be using student laptops in the
regular laboratory to run the program, if enough students can bring laptops to class for this to be feasible.
This will be discussed during the first week of class. Although you can log onto the network and use the
Neurosim outside of the computer lab, all students should attend the laboratory session for instruction on
how to work with the program and to find out what portions of the program will be used for the
laboratory report. Specific instructions for the experiments required for class are given below. You will be
shown how to load parameter files for each set of experiments, to allow you to focus on specific aspects
of neurophysiology. To obtain context-specific help for the Neurosim program, either press the F1
function key, or select "help" from the menu bar at the top of the screen.
EXPERIMENTS TO PERFORM
Students may work either alone or in pairs to perform the experiments. The experiments which you
should complete in the lab period are outlined below. A report describing these experiments will be due in
lab next week. You may do additional experiments during open lab hours, and turn in a report for up to 15
points of extra credit. Additional details are provided below (see "Extra Credit", p. 69). The due date for
extra credit will be announced in class.
62
LABORATORY REPORT
A general description of the style and format for the laboratory report is given at the beginning of the lab manual. In
brief, your written report should include 1) a title, 2) an introduction, which states the hypotheses tested in your
own words; 3) a methods section, explaining how you set up the stimulator and oscilloscope, and what ionic
conditions were used for your experiments (you don’t need to explain exactly how you started the computer
program, but should clearly explain the “experimental” conditions you used); 4) a results section, summarizing the
data collected to test these hypotheses, in tabular form if appropriate; and 5) a discussion section, describing the
conclusions that can be drawn from the data collected and supporting these conclusions with specific data from the
results section. Data from your experiments can be saved by using the "measure" option available on the "results"
screen. Traces of the oscilloscope screens to accompany your report may be made by first choosing the file, print
setup option, followed by the file, print option; alternatively, you can click on the printer icon on the menu bar.
Your report should include any calculations used for interpretation of results. Remember that all conclusions should
be supported by specific data. The report should be written as an integrated description of all the experiments
outlined below. That is, there should be a single introduction, a single results section, and a single discussion
section. Within each section, make clear and logical transitions between the different experiments described. You
may find it useful to use section headings to separate the different experiments, but do not prepare sequential lab
reports describing the different experiments. Instead, your report should show how these experiments are related.
There are many ways to save data to use in your report. Ask your instructor if you are not sure how to save or
print information. It is your responsibility to save data in a form that can be used to prepare the laboratory
report.
Key information to include in laboratory reports (see sections below for more detail)
K
H
p
Measurements showing a threshold stimulus for an action potential.
o A range of values should be included, demonstrating that there is a threshold; reporting a single
value is not sufficient.
o As you test a range of stimuli, you should make qualitative observations – did an action potential
occur or not? If an action potential occurred you can make quantitative observations – did the
intensity of the stimulus affect the characteristics of the action potential?
o A table showing a range of values tested should be included. Although it is quite reasonable to
jump around while initially testing different stimuli, the values in this table should be arranged in a
clear and logical order.
o Oscilloscope tracings can be very useful to clarify what is meant by an action potential occurring.
However, you need not include a tracing for every stimulus tested.
Measurements demonstrating a refractory period between stimuli
o A range of values for delay between the two stimuli is needed to show that there is a refractory
period; a single value is not sufficient.
o You should observe whether delays longer than the minimum required for a second action
potential to occur affect the characteristics of the action potential.
o The delay between stimuli is the time from the end of one stimulus until the beginning of the next
stimulus. Since the first stimulus takes some time to elapse, you will need to account for this time
period when calculating the delay and reporting the refractory period.
Using voltage clamping to determine the internal sodium concentration in a neuron
o A range of voltage clamp values should be tested, paired with observations of the Na+ current.
Your objective is to determine the membrane potential that gives a Na+ current as close to zero as
possible.
o Oscilloscope tracings with positive, negative, and minimal Na+ currents will be useful to
demonstrate this concept.
o A graph of the Na + currents observed as a function of the clamp potential voltages tested should
be included.
o Calculations to determine the internal Na+ concentration should be included in the discussion
section.
63
PART ONE: All-or-none properties of the action potential; threshold stimulus for the action
potential.
One of the important properties of the action potential is the
"all-or-none" phenomenon, described in lecture and your
text. This experiment will be run in "current clamp" mode,
in which the stimulator will give a single pulse of current to
the nerve. You will then observe changes in the membrane
potential as measured on the oscilloscope screen.
Load the parameter file BI313_1.hh by selecting File-Open
from the menu bar. [This file is in the
Lrc-class\INSTRUCT\Biology\jrebers\Neurowin\Hh
directory; see me for help if the BI313_1.hh file does not
appear on the list of files to load.] Click on the results
screen visible to the left to see the stimulator settings and
the ionic conditions for the nerve. Press the start button to
initiate a stimulus. On the results screen, the upper trace
will indicate the membrane potential for the nerve (in units
of millivolts, or mV), and the lower trace will show a trace
of the stimulus (in microamperes, or µA). The horizontal
axis for each scale gives the time elapsed in milliseconds
(ms). The parameters are initially set to give a 50 µA
stimulus with a duration of 0.25 ms. A stimulus of this
magnitude will give an action potential, with the
Figure 1. Oscilloscope tracings after a 50 µA, 0.25 ms
oscilloscope traces looking like those in Figure 1.
stimulus.
The initial membrane potential of the nerve before the stimulus is known as the resting potential. After the
stimulus, the membrane depolarizes, becoming less negatively charged inside the cell, reaching a peak
where the membrane potential is positive. The membrane then undergoes a transient hyperpolarization
(becomes more negative than the resting potential).
You can measure the value of the membrane potential at a particular time by using the measure option on
the results screen. Press the measure button, and a table will appear on the left of your screen. The
"Trace" box at the bottom of the table should read "voltage". Move the mouse cursor over the
oscilloscope trace - as you do so, the arrow will change to crosshairs. Position the crosshairs over the
membrane potential trace prior to the stimulus, and press the left mouse button. The membrane potential
and time it was measured will appear in the table in the measure dialogue box. The first column contains a
simple numerical identifying the order in which the measurements are made. One way to make this more
informative is to click on the box labeled "User", and type in "Stim - uA" for stimulus, including the
units. Then move the cursor to the first row under stimulus, click, and enter 50 for the 50 µA stimulus.
Repeat the measurement at the peak of the action potential. Relabel the "stim" to indicate that another 50
µA stimulus was given.
Now click on the left hand side of the screen to switch to the setup display. Under the current clamp
settings, double click on the amplitude of pulse 1 (amp), which is currently set to 50 µA. Type in 5 to
change the amplitude of the stimulus. Press the start button to initiate a new stimulus. A second trace will
appear on the results screen, which does not show the depolarization characteristic of an action potential.
64
Repeat your measurements with a variety of stimuli, keeping track of whether or not an action potential
occurs. The minimum stimulus needed to cause an action potential is known as the threshold. Note that
for values below the threshold, there may be a small membrane depolarization, but the large spike
depolarization does not appear. Why do you think this difference is observed?
If an action potential is observed for a particular stimulus, record the value of the stimulus, the membrane
potential at the peak of the action potential, and the time at which the peak of the action potential is
observed. The methods section for your report should provide enough information so the "experiment"
can be replicated. In particular, you need to note the stimulator settings used and provide the internal and
external ionic concentrations for the neuron. The results section should include a table showing the
measurements you made, labeled clearly with all appropriate units included. A figure showing a trace of
the oscilloscope readings from a typical experiment would also be useful. (You should not include
individual traces for all stimuli however - these values are more easily displayed in tabular form.) In the
discussion section of your report, you should address the following questions. Do either the peak voltage
of the action potential or the time the action potential reaches the peak change for different stimuli above
threshold? If so, what kind of relationship is observed between stimulus strength and these variables?
How do your observations compare to previous experiments done with squid axons?
Technical notes about measurements:
The values from the table in the "measurement view" can be copied to the Windows clipboard, using
the copy button. You can then switch to a word processing program or to the Notepad program in
Windows to paste these values into a document. The values can then be saved for your report.
Alternatively, you may find it simpler to record the values using pen and paper. Either way is
acceptable, providing that your report provides a clear and organized table of results.
The results view records the results of successive stimuli on the screen. To clear the screen, use the
clear button to the left.
For some measurements, you may find it useful to change the horizontal or vertical axes on the
oscilloscope screen. First, clear the screen of previous readings. Then double click on the value you
wish to change (for example, the 10 millisecond value on the horizontal time axis. When the number
is highlighted in blue, enter the new value.
The View-Datum lines option can be used to add horizontal or vertical lines to the traces to assist in
alignment of different experiments. If you want to get fancy, the View-Trace ID and ViewAnnotation options can be used to add labels to your graphs. See the on-line help manual for details.
Before moving on to part two, you may find it useful to make some observations using two additional
traces on the oscilloscope. Clear any previous experiments, and then choose View-Trace Display. Select
the boxes next to "Current-Show Graph" and "Conductance-Show Graph". Press the OK button at the
bottom, and then press the Start button to initiate a stimulus. Two new traces will be displayed,
indicating the changes in membrane current and the changes in membrane conductance as the action
potential occurs. Measurements and discussion of these changes are not required for your lab report.
However, comparing the changes in membrane current and conductance to the changes in membrane
potential will help you to understand how ion channels in the axon membrane give rise to an action
potential.
65
PART TWO: Refractory period
Reload the parameter file BI313_1.hh by selecting File-Open from the menu bar, to return all conditions
to the original values. Set the value for the amplitude of
pulse 2 to 50 µA, to match the value for pulse 1, leaving the
other settings unchanged. Select start to initiate a stimulus,
and observe the action potential. You should see something
like Figure 2. Note that the first stimulus pulse causes an
action potential but that the second does not, even though
both stimuli are the same strength.
Press clear on the results screen, double click on the 10 at
the right-hand side of the horizontal time scale, and change
the value to 30. Now choose Window-Setup, and change
the delay for pulse 2 to 20 ms. Press start to initiate a
stimulus. Note that two action potentials are now observed.
Note: it is important to rescale the time axis so you can see
both stimuli after increasing the delay between them. You
may find it helpful to change the scale on the time axis
again for more precise measurements.
During an action potential, voltage-gated Na+ channels
open in response to a membrane depolarization, and then
change to a closed and inactive state. The time for the
voltage-gated Na+ channels to recover so they will once
Figure 2. Oscilloscope trace after two 0.25 ms, 50 µA
again open in response to membrane depolarization is
stimuli, with 7 ms between stimuli.
known as the refractory period. You can measure the
length of the refractory period by changing the delay between the two stimuli, and observing whether or
not an action potential occurs in response to the second stimulus. The minimum time delay between the
two stimuli for the production of a second action potential is the refractory period. Your lab report should
include the measurements you made to determine the refractory period, and the discussion should relate
these measurements to what you know about the properties of the axon membrane. Provide a data table
with a range of measurements, including delays between pulses shorter than the refractory period and
longer than the refractory period. Do delays longer than the minimum affect the characteristics of the
action potential?
PART THREE: USING THE VOLTAGE CLAMP TECHNIQUE TO DETERMINE
INTRACELLULAR ION CONCENTRATIONS
BACKGROUND - VOLTAGE CLAMPS AND THE NERNST EQUATION
The experiments you have done so far measured the change in membrane potential in response to a pulse
of current that caused different ion channels in the membrane to open and close in succession. These
changes in channel properties result in changes in membrane conductance; the membrane potential then
changes as ions flow across the membrane. One challenge in investigating the changes in membrane
properties is that the channel properties, and hence the membrane conductance, are voltage-dependent.
That is, as the membrane potential changes, ion channels open and close, and so the membrane
conductance changes with time. As the conductances change, the currents flowing through the channels
change! There is an intricate series of feedback loops regulating the properties of the membrane at a
specific instant.
66
A voltage clamp is an electronic device that provides a way to break these feedback loops, setting the
membrane potential at a level fixed by the user. The circuit detects any deviation of the current from the
set value, injecting current to counteract any deviation. The amount of current that must be injected is
exactly equal and opposite to the ionic current flowing through the ion channels in the membrane.
In order to use voltage clamping to determine the intracellular ion concentrations, you will need to make
use of the Nernst equation, which gives the relationship between the equilibrium potential for an ion and
the ratio of intracellular and extracellular concentrations of that ion.
E
E
RT ln [ion] ext
zF
[ion] int
equilibrium potential (in volts) for a particular ion; the membrane potential required to prevent net
movement of that ion across the membrane at the internal and external ion concentrations specified [ion]ext and [ion]int
R Universal gas constant (2 cal mol-1 ºK-1)
T
Temperature, in degrees Kelvin
z
valence of the ion of interest
F
Faraday's constant, an electrical constant (2.3 x 104 cal V-1)
For Na+ ions, with a valence of +1, and a temperature of 6ºC, the Nernst equation can be simplified to:
EmV
24 ln
[Na ext]
[Na int]
If you know the internal or external concentrations of an ion, you can use the Nernst equation to calculate
the amount of voltage required to prevent ion flow. As we will discuss in lecture, when the membrane is
at rest, Na+ channels are closed. When the channels open in response to a membrane depolarization, Na+
ions flow out of the neuron until the sodium equilibrium potential is reached. The equilibrium potential is
the membrane potential at which the direction of Na+ ion flow through the membrane will be reversed.
The direct measurement of the reversal potential can be done with a voltage clamp. Using this value, it is
then possible to determine what the internal concentration of Na+ ions inside the neuron is, provided that
the external concentration is known. In order to limit current flow through the membrane, you will add
drugs to block some of the ion channels. Tetrodotoxin, or TTX, is a drug isolated from puffer fish that
will block voltage-gated Na+ channels. Tetrethylammonium, or TEA, blocks voltage-dependent K+
channels. Scorpion toxin prevents inactivation of the voltage sensitive Na+ channels (there are a number
of other toxins found in scorpion venom which we won't be concerned with).
67
PROCEDURE - MEASURING [Na+] USING VOLTAGE CLAMPING
Load the parameter file BI313_2.hh by selecting File-Open
from the menu bar. Make sure the mode in setup view is set to
voltage clamp, and that the values listed under the voltage
clamp are: Hold, -70 mV, 0.5 ms; Clamp 1, 0 mV, 5.0 ms,
Clamp 2, 0 mV, 0 ms. Click start to initiate a stimulus. You
should see a trace like the line in the bottom panel of Figure 3
labeled "Total current", showing the net value of all ion flow
through the membrane. In order to measure the Na+
equilibrium potential, we need to study the Na+ current in
isolation. Although we can't measure Na+ current specifically,
we can block K+ channels, providing a view of membrane
current due only to the Na+ channels. To do this, check the box
next to TEA in the "drugs" box to inactivate the voltage-gated
K+ channels. Now you should see both lines shown on Fig. 3.
Clear the screen in results view, and repeat the experiment,
changing the Clamp Potential parameter for Clamp 1 (adjusted
in setup view) from 0 to + 60 mV in 10 mV steps. If the
voltage clamp holds the membrane potential to a value of 0
mV, the initial flow of Na+ ions will not be affected by a
voltage gradient, so Na+ ions will flow from high
concentration outside the cell to low concentrations inside the
Figure 3. Voltage and current traces in voltage clamp
cell, expressed by convention as a negative Na+ current, as
experiment.
seen in Figure 3. As the membrane potential is made more
positive with the voltage clamp, the positive charge inside the cell opposes the chemical gradient. For
membrane potentials as high as +60 mV, the flow of Na+ ions will actually be from inside the cell to the
outside, since the electrical gradient favors an outward movement of positively charged ions. This is
indicated by the positive Na+ current seen in Figure 3. As you increase the voltage from 0 to + 60 mV,
you should notice a transition point at which the initial Na+ current is close to 0. This is the equilibrium
potential, the value for the membrane potential where the electrical gradient of charge across the
membrane exactly balances the chemical gradient of Na+ ions on either side of the membrane.
The data collected for this section of the lab should include a graph showing the clamp potentials ranging
from 0 to +60 mV and the corresponding plots of Na+ current. When you find a voltage clamp potential
close to the equilibrium potential, it will be helpful to increase the resolution of the display. Clear the
display, and then click the lower scale on the current display and set it to -100; then click the upper scale
and set it to +100. Repeat your measurements with a range of values for the voltage clamp that will allow
more precise measurement of the value for the Na+ equilibrium potential. You will find it helpful to use
the measure option on the results screen to get precise values for the Na+ current for the different voltage
clamp values. Use the simplified form of the Nernst equation for Na+ ions at 6ºC to determine the internal
Na+ concentration for the neuron. Since you just determined EmV, the Na+ equilibrium potential
experimentally, and the [Na+ext] is shown on the Setup screen, calculating the [Na+int] is a straightforward
algebraic calculation. See the appendix to this lab on using exponents and logarithms if you need a review
of these topics. Be sure to show your work for all calculations.
68
Warnings
The authors of this program know a lot more about neurophysiology than your instructor. I have
selected specific aspects to illustrate some basic ideas about nerve cell function. If you delve deeply
into the program, you will find it easy to stump your instructor. I will help as much as I can, but can
make no guarantees of explaining everything you encounter. Lab reports and extra credit must be
clearly written to show that you fully understood any experiments you did.
These programs are based on mathematical models of the nerve cell, based in large part upon the
work published by Hodgkin and Huxley. Probably the greatest problem in using a model is that it
works every time, without the challenges and frustrations encountered when working with living
cells.
The programs are set up to limit input conditions to a biologically reasonable range. It may be
possible, however, to discover some combination of conditions that gives peculiar results because of
limitations of the model rather than because of the biology of the system.
EXTRA CREDIT
A few suggestions for extra credit experiments are outlined below. For other possibilities, see Panel 11-3
in MBOC (p. 531) or the Scientific American articles listed at the beginning of the handout. A maximum
of 15 points will be awarded for extra credit. For complete credit, the experimental hypotheses to be
tested must be explained clearly, data presented in an organized and concise fashion, and the way in
which the data supports your hypotheses should be explained clearly. The number of points awarded will
depend upon how thorough the experiments are and how clearly they are described.
You will have the greatest flexibility for your experiments if you exit the program using File-Exit or the
check box in the upper right, and restart the program. This will allow you to view the full range of
parameters that can be adjusted. Help for setting up the experiments or determining allowable parameters
is available from the help menu, which has a searchable index and a table of contents.
The most abundant ions inside and outside the axon are Na+, K+, and Cl-. The [Na+] and [K+] inside
and outside of the axon can be manipulated from the setup view. Test the effect of changing the ionic
conditions upon both the resting membrane potential and the peak height of the action potential. The
acceptable range for ion concentrations is 0.01 to 1000 millimoles per liter. This experiment is best
done using current clamp mode.
Temperature can be varied from 0 to 30 degrees Centigrade. Test the affect of changing the
temperature upon the values for threshold stimulus or peak height for the action potential (other
values can be tested - these are simply suggestions).
Test to see whether the nerve can initiate an action potential in the presence of the three different
toxins available. It may be of interest to record the ion currents and membrane conductance at the
same time. Explain the effects you observe based upon what you know about ion channels and the
action potential.
69
APPENDIX - REVIEW OF EXPONENTS AND LOGARITHMS
In order to calculate the internal concentration of an ion when the external concentration and equilibrium
potential for that ion are known, you will need to do some algebra using exponents and logarithms. You
may find the relationships below useful, as a review of how to work with these functions. Remember that
logarithms are most commonly expressed in either base 10 [log10 (x)] or in the base of natural logarithms
(e), as ln (x).
Equations can be expressed using either exponents or logarithms. For example:
A = log10 B
is equivalent to:
10A = B
Or, for natural logarithms:
A = ln B
is equivalent to:
eA = B
When solving equations, you may find the relationships below useful.
Laws of Exponents
Laws of Logarithms
am an = am+n
ln (xy) = ln(x) + ln(y)
(am)n = amn
l xl
ln ll ll l ln(x) - ln(y)
l yl
(ab)m = am bm
a
b
m
a -m a
ln x n n n ln(x)
m
a
bm
ln(e) = 1; log10 (10) = 1
or generally, loga (a) = 1
1
am
ln (1) = 0; log10 (1) = 0,
or generally, loga (1) = 0
am
a a m -n
n
a
Information on exponents and logarithms adapted from College Algebra and Trignometry, B. Kolman &
A. Shapiro (1986).
70
LAB CRITIQUE SHEET
Laboratory 5
Neurophysiology
Clarity of written introduction/background
Length of written introduction/background
Clarity of in-lab introduction/background
Length of in-lab introduction/background
Clarity of procedures
1 2
Unclear
3
4 5
Very
Clear
2
3
4 5
Too long
1 2
Unclear
3
4 5
Very
Clear
2
3
4 5
Too long
1 2
Unclear
3
4 5
Very
Clear
1 2
Low
3
4
5
High
1
Too short
1
Too short
Relevance to lecture material
Rate this lab relative to
other BI 313 labs
Rate this lab relative to
other labs at NMU
1
Poor
2
3
4
5
Excellent
1
Poor
2
3
4
5
Excellent
As new labs are introduced, some labs may be deleted. Is this lab a keeper?
Specific errors in lab manual, or clarifications needed.
Other comments. (Use back if needed)
71
Comments
72
LABORATORY 7
SPECTROPHOTOMETRIC ASSAY OF MITOCHONDRIAL SUCCINATE
DEHYDROGENASE
Background Reading (required):
Molecular Biology of the Cell, pp 128-135; 162-165; pp 653-667
Background Reading (optional):
Hinkle, P.C. and McCarty, R.E. (1978) "How Cells Make ATP", Scientific American, 238 #4, pp
104-123.
Although this article may be older than you are, the ideas presented about ATP synthesis
are still valid and are clearly presented.
http://www.nobel.se/chemistry/laureates/1997/press.html
Press release from the Royal Swedish Academy of Sciences, upon the award of the 1997
Nobel prize in Chemistry to Professor Paul D. Boyer, University of California, Los
Angeles, USA, and Dr. John E. Walker, Medical Research Council Laboratory of
Molecular Biology, Cambridge, United Kingdom for their elucidation of the enzymatic
mechanism underlying the synthesis of adenosine triphosphate (ATP). The press release
gives a concise description of the mechanism used by ATP synthase to make ATP in
mitochondria.
IMPORTANT SAFETY NOTE
We will be using sodium azide to inhibit electron transport in cauliflower mitochondria in this lab. This
compound is equally effective at inhibiting electron transport in Homo sapiens.
1) Do not mouth pipet anything in the lab. Use the pipeting devices provided.
2) No eating or drinking is permitted in the lab. If you have an empty pop can, please keep it on the
floor, so it will be clear that you are not drinking from it.
INTRODUCTION
In this laboratory we will first isolate mitochondria from cauliflower using a procedure called cell
fractionation, and then study the activity of succinate dehydrogenase, one of the enzymes found in the
mitochondria. Cell fractionation is a procedure used by cell biologists to study the function of different
parts of the cell in isolation. The cells are first disrupted, or homogenized, and then physical differences
between the cellular components are used to separate them. One common tool for separating the different
parts of the cell is the centrifuge. Large organelles, like nuclei or chloroplasts, can be separated using the
relatively low forces that can be generated by a table-top centrifuge. Smaller organelles, like
mitochondria, require a high-speed floor model centrifuge. An ultracentrifuge, which can generate forces
more than 100,000 times that of gravity, can be used to separate small organelles, viruses, and even large
macromolecules.
73
HOMOGENIZATION
Before the different organelles can be separated from a cell, the plasma membrane must be broken open
without disrupting the organelles. In the case of plant cells, the cell wall, which is much stronger than the
plasma membrane, must also be disrupted. Treating cells with detergents to disrupt the plasma membrane
would not work, since detergents would disrupt internal membranes as well as the plasma membrane.
Some of the techniques which have been used to disrupt cells are grinding in a mortar, using a blender
(sometimes the same as a household blender, sometimes a fancier model designed for cells), ultrasonic
vibration, or osmotic shock. The type of buffer used for homogenizing the cells can affect the recovery of
intact organelles. After homogenization, the plasma membrane and the endoplasmic reticulum are
disrupted and reseal to form small vesicles called microsomes, but the rest of the organelles can be
recovered intact. Homogenization is usually done at 4ºC to prevent proteins from denaturing and to
inhibit the action of degradative enzymes.
CENTRIFUGATION
The organelles in the cell differ in their size, shape, and buoyant density. After disrupting the cells and
making a disorganized stew of the cellular components, the organelles can be separated by spinning the
homogenate at relatively high speeds.
The type of centrifugation we will carry out in lab is known as velocity sedimentation (or as rate-zonal or
differential-velocity centrifugation). This procedure separates the organelles on the basis of size and
density; larger organelles are subjected to the greatest centrifugal force and sediment more rapidly than
smaller organelles. As shown in Figure 4-35, MBOC, a low speed centrifugation [1,000 times the force of
gravity (1,000 g), for 10 minutes] will separate whole cells, nuclei, and chloroplasts. A medium-speed
spin (20,000 g for 20 min) will separate mitochondria, lysosomes, and peroxisomes. A high-speed spin
(80,000 g for 1 h) will separate microsomes and small vesicles, and a very-high speed spin (150,000 g for
3 h) will sediment ribosomes, viruses, and large macromolecules.
The fractions obtained by rate-zonal centrifugation are usually not completely pure. For instance,
lysosomes, mitochondria, and peroxisomes will often be found in the same fraction. Equilibrium densitygradient centrifugation can be used if fractions of higher purity are required. In this procedure, the
separation is based on density alone, rather than on a combination of size and density. The fraction to be
purified is layered on top of a gradient of a dense nonionic solution made up of glycerol or sucrose. The
tube is centrifuged at very high speed for several hours, so that the different particles migrate to a position
such that their density is equal to that of the surrounding medium and cannot sediment any further. (See
Fig. 4-36, MBOC).
The force generated by a centrifuge depends not only on the angular velocity of the rotor, but also upon
the distance of the sample from the center of rotation. For this reason, methods sections in scientific
papers usually report the relative centrifugal force (RCF) in g units as above, where one g is the normal
force of the earth's gravity. For a given angular velocity, different rotors will exert different RCFs upon a
sample. The relative centrifugal force can be calculated using the equation below, where rpm is the
angular velocity of the rotor in revolutions per minute, and r is the distance in centimeters from the center
of rotation to the sample.
RCF
1.119 x 10
5
r rpm
2
Alternatively, a table or graph is often provided with a centrifuge rotor to convert a given rpm value to the
RCF in gs.
74
The purity of the different organelle fractions can be assessed either by examining the different fractions
with a microscope, or by assaying marker enzymes or proteins that are characteristic of particular
organelles. Some typical markers are listed in Table 1.
Table 1. Markers for different organelles.
Organelle
Marker
Mitochondria
Cytochrome C
Succinate Dehydrogenase
Peroxisomes
Catalase
Lysosomes
Acid Phosphatase
Golgi Apparatus
Glycosyl Transferase
ELECTRON TRANSPORT IN THE MITOCHONDRION
The role of the mitochondrion is to produce ATP by oxidizing reduced hydrocarbons. Catabolic reactions
that begin in the cytoplasm are carried to completion in the mitochondrion. Oxidative metabolism in the
mitochondrion begins with acetyl CoA, produced either from pyruvate or from fatty acid metabolism. The
acetyl CoA (2 C) combines with oxaloacetic acid (4 C) in the matrix to produce citric acid (6 C). A series
of oxidation/reduction reactions, known as the citric acid cycle, tricarboxylic acid (TCA) cycle, or Krebs
cycle, produces NADH, GTP, FADH2, and CO2, and regenerates the oxaloacetic acid (See Fig. 14-14,
MBOC). Two important features of the citric acid cycle to remember are 1) oxygen is not directly
consumed; and 2) ATP is not directly produced. Although the molecule of GTP produced can be readily
converted to ATP by an exchange reaction with ADP, most of the ATP produced in the mitochondria
comes from a separate series of electron-transport reactions coupled to proton pumping from the matrix to
the intermembrane space. ATP synthetase bound to the inner membrane uses the energy stored in the
electrochemical proton gradient formed across the mitochondrial inner membrane to synthesize ATP in
the matrix.
The citric acid cycle transfers electrons from acetyl CoA to NADH and FADH2 in a series of oxidationreduction reactions. One of the enzymes required is
HO
O
HO
O
succinate dehydrogenase (SDH). Succinate
C
C
dehydrogenase catalyzes the dehydrogenation
(oxidation) of succinate to produce fumarate (see Fig.
CH
CH2
1). The electrons removed from succinate are
CH
CH2
transferred to FAD to produce FADH2. The FAD is
covalently bound to the SDH molecule. Since FADH2
C
C
is bound to protein, it is not free to diffuse in the
HO O
HO O
matrix to transfer electrons, like NADH does.
Instead, the FADH2 is reoxidized by transferring its
Fumarate
Succinate
electrons to the electron transport chain on the inner
membrane of the mitochondrion. This transfer is
facilitated by the fact that SDH, unlike other enzymes Figure 1 Dehydrogenation (oxidation)
in the citric acid cycle, is bound to the inner
of succinate produces fumarate.
membrane instead of being found in the matrix. The
75
electrons are transferred from FADH2 to ubiquinone (coenzyme Q), one of the carriers in the electron
transport chain (see Fig. 14-33, MBOC). The electrons in FADH2 do not have quite as much energy as do
those in NADH. The transfer to ubiquinone bypasses NADH dehydrogenase, the first respiratory enzyme
complex in the electron transport chain, so fewer protons can be pumped across the mitochondrial inner
membrane when FADH2 is oxidized than when NADH is oxidized.
MEASUREMENT OF SUCCINATE DEHYDROGENASE ACTIVITY
One way to follow the reaction shown in Fig. 1 would be to measure the disappearance of succinate or the
appearance of fumarate after adding succinate to isolated mitochondria. This could be done with great
precision by using radioactively labeled succinate, but the reagents required are expensive and potentially
hazardous. We will use a less direct procedure that is simpler to work with in the lab. An artificial
electron acceptor will be added to a reaction mixture that contains mitochondria and succinate. The
electrons removed from succinate by SDH will be transferred to the artificial electron acceptor, and the
reduction of this acceptor will be measured. In order for the artificial electron acceptor to work, the
normal electron transport pathway must be inhibited. This will be done by adding sodium azide (NaN3) to
the reaction mixture. This substance binds tightly to the cytochrome oxidase complex in the inner
membrane of the mitochondrion and inhibits the electron transport chain. Cyanide and carbon monoxide
have the same effect upon cytochrome oxidase.
The artificial electron acceptor we will use to monitor SDH activity is 2,6-dichlorophenol-indophenol
(DCIP). The oxidized form of DCIP is blue and the reduced form is colorless. As electrons are removed
from succinate and transferred to DCIP by SDH, the dye will change from blue to clear. The reaction can
be summarized as:
SDH - FADH2 DCIP oxidized (blue)
SDH - FAD DCIP reduced (colorless)
In
this assay, you will measure the disappearance of a colored substrate. In other spectrophotometric assays,
the appearance of a colored product is measured. The oxidized form of DCIP strongly absorbs light at the
red end of the spectrum. We will measure the absorbance of the reaction mixture when light with a
wavelength of 600 nm is passed through the mixture. This will give us a sensitive and quantitative way to
measure the amount of DCIP that remains in the reaction mixture.
ENZYME KINETICS
Not all enzymes work with equal efficiency. The properties of a particular enzyme can be revealed by
studying its kinetics - the reaction rate for that enzyme at different enzyme and substrate concentrations.
According to a model proposed by Leonor Michaelis and Maud Menton in 1913,
E S
k1
ES
k3
E P
k2
An enzyme, E, combines with a substrate, S, to form an enzyme-substrate complex. This complex forms
at rate k1 and breaks down to form E + S at rate k2. The enzyme-substrate complex can also form a
product, P, at rate k3. At the initial stages of the reaction, the concentration of the product will be so low
that the rate of conversion of E + P to ES will be negligible.
Rate constants k1, k2, and k3 can be combined as a new constant called the Michaelis constant (KM).
76
k2 k3
k1
KM
The rate of reaction for a particular enzyme can be expressed using the Michaelis-Menten equation:
V
Vmax
[S]
[S] KM
At very low substrate concentrations ( [S]<<KM ), the rate (V) will be directly proportional to the
substrate concentration. At high substrate concentrations, all of the enzyme molecules will be saturated
with substrate, and the reaction rate will be at a maximum (Vmax) for that concentration of enzyme. When
[S] = KM, V = Vmax/2; that is, the reaction will proceed at half the maximum velocity when the substrate
concentration is equal to KM. Thus, KM can be directly determined by measuring the reaction rate at
different substrate concentrations.
Another form of the Michaelis-Menten equation is:
V0
k3 [E][S]
k2 k3
[S]
k1
Where Vo is the initial velocity of the reaction; that is the reaction rate immediately after enzyme and
substrate are mixed together. This form of the Michaelis-Menten equation emphasizes the fact that
the initial velocity depends on both enzyme and substrate concentration. This is one of the key
concepts you will test in this laboratory.
In your experiments, you will measure the initial velocity of the succinate → fumarate reaction using
several concentrations of succinate dehydrogenase. To vary the enzyme concentration, you will add
different volumes of resuspended mitochondria to the reaction mixture. The total reaction volume and
substrate concentration will be held constant, and the relative volume of mitochondria will be a measure
of the relative concentration of SDH added to the reaction. The concentration of substrate (succinate) will
be high enough so that in the initial stages of the reaction all enzyme molecules will be saturated with
substrate and Vo should be directly proportional to enzyme concentration.
The rate of reaction will be measured by monitoring the change in absorbance (ΔA600) with time. As the
reaction proceeds, the DCIP will be reduced, and change from blue to colorless. The rate of that change is
directly proportional to Vo. You will plot ΔA600 vs. time for three different enzyme concentrations. For
each plot the initial slope of the plot can be used as a measure of the rate of change of A600. The initial
velocities determined from these plots will then be graphed as a function of enzyme concentration.
The instrument we will use to measure the ΔA600 is a spectrophotometer. Light of defined wavelength
passes through a sample and strikes a photocell. The photocell then gives an electronic reading either of
the fraction of light that has passed through the sample (% transmittance, T) or the amount of light
absorbed by the sample (absorbance, A). The instructions on p. 82 explain how to use the Milton Roy
Spectronic® 20 spectrophotometer.
COMPETITIVE INHIBITION
77
Enzymes can be inhibited by specific small molecules or ions. In some cases, this inhibition represents an
important regulatory step — for example, the reaction product of many enzymes can inhibit more product
from being formed (feedback inhibition). In other cases,
drugs or toxic agents may act by inhibiting specific
HO
O
enzymes. An irreversible inhibitor binds tightly to the
C
O
OH
enzyme and will not dissociate. A reversible inhibitor, on
CH2
the other hand, will dissociate rapidly after forming an
enzyme-inhibitor complex. The extent of inhibition
CH2
CH2
depends on the kinetics of binding inhibitor, just as the
C
C
extent of reaction depends on the extent of binding
substrate.
HO O
HO O
In some cases, molecules inhibit enzymatic reactions
Malonate
Succinate
because they compete with substrate for binding to the
active site of the enzyme. Malonate, which is one of the
Figure 2 Succinate and malonate
reactants in the citric acid cycle, can act as a competitive
have related structures.
inhibitor for SDH. As shown in Fig. 2, the structures of
malonate and succinate are quite similar. Malonate can
bind to the active site of SDH, but does not have hydrogen atoms in the proper configuration to be
dehydrogenated by the enzyme. If the amount of malonate is high relative to the amount of succinate,
most molecules of SDH will have malonate bound and will be unable to pass electrons on to DCIP. When
enzyme velocity is reduced due to the presence of a second molecule that can compet for binding to the
enzyme , an effect known as competitive inhibition is observed.
CONTROLS
In one of the reaction tubes in Table 1 enzymes will be inactivated by heating the mitochondrial
suspension. This will test the hypothesis that the color change is due to active enzymes in the
mitochondria. There are several other controls for the assay that you can design to include in your
experiment. Some suggestions are given below.
ISOLATION OF CAULIFLOWER MITOCHONDRIA
We will isolate mitochondria from cauliflower florets by differential centrifugation. Each group of three
students should prepare a cauliflower homogenate and isolate mitochondria as directed below. First, the
cauliflower will be ground in a mortar with a buffered, isotonic mannitol solution, using sand to help
break open the cell walls. The homogenate will be filtered through cheesecloth to remove the larger
pieces of cauliflower, and then centrifuged for 10 minutes at 600 g to remove nuclei, unbroken cells, and
sand. The supernatant will then be centrifuged for 30 minutes at 10,000 g to sediment the mitochondria.
All solutions and equipment used for the mitochondria isolation should be kept ice-cold, to prevent
denaturation of the succinate dehydrogenase and to keep the mitochondria intact.
78
PROCEDURES
ISOLATION OF MITOCHONDRIA FROM CAULIFLOWER (Brassica oleracea)
1. Use a razor blade to shave off the outer 2-3 mm from cauliflower florets. Each group of three students
will need 20 g of cauliflower shavings. Collect the shavings in a petri dish until ready for step 2.
2. Place the cauliflower shavings in a large chilled mortar with 40 ml of ice-cold mannitol grinding
medium and 5 g of purified sand. Grind the tissue with a chilled pestle for 4 minutes.
3. Filter the homogenate through 4 layers of cheesecloth into a chilled 50 ml centrifuge tube. Do this by
placing a funnel in the centrifuge tube and lining the funnel with the cheesecloth. Wring any residual
juice from the cheesecloth.
4. Bring your tube to the ice bucket on the front bench.
5. Balance your tube of cauliflower homogenate with a tube from another group. Centrifuge the tubes at
1600 rpm (600 g) at 4ºC for 10 minutes.
6. Carefully decant the supernatant into a clean, chilled 50 ml centrifuge tube.
7. Centrifuge the post-nuclear supernatant at 12,000 rpm (10,000 g) at 4ºC for 30 minutes.
While the mitochondria are spinning, you can begin preparing your tubes for the
succinate dehydrogenase assay, as directed below.
8. Discard the supernatant and add 15.0 ml of ice-cold mannitol assay medium to the pellet.
9. Use a spatula to dislodge the pellet from the wall of the tube, and then use a Pasteur pipet to suspend
the mitochondria. Be sure that the no clumps remain, since these would cause variation in the amount
of enzyme in different tubes.
10. Save the tube containing the resuspended mitochondria on ice until ready for the SDH assay.
79
MEASUREMENT OF SUCCINATE DEHYDROGENASE ACTIVITY
BASIC ASSAY
The procedures below give the basic assay procedure for measuring succinate dehydrogenase activity, and
explain how to measure the effect of changing enzyme concentrations upon initial velocity of the enzyme
reaction. Important: you will need to plan additional assays to ask experimental questions, as
described below. You should do this before coming to the lab.
1. Turn on the spectrophotometer and allow it to warm up at least 5 minutes. Set the wavelength to 600
nm. Complete instructions are given on p. 82.
2. Prepare a boiling water bath using a beaker with boiling chips and a hot plate. The boiling water bath
should be kept at a low simmer until the mitochondria are ready, and then turned up to a full boil for
step 4.
3. Dispense a 0.8 ml aliquot of the mitochondrial suspension to a 16 x 100 glass tube. Heat this tube in
the boiling water bath for 5 minutes, and cool the tube in an ice-water bath.
4. Label 7 cuvets as shown in Table 1 (p. 81). Arrange the cuvets in a plastic test tube rack, to avoid
scratching.
5. Follow the directions in Table 1 to add assay reagents to the tubes as directed. Do not add the
mitochondrial suspension at this time. All solutions should be at room temperature, except the
mitochondrial suspension, which should be kept on ice. Keep careful track of the reagents added,
since not every tube receives the same components. You may find it helpful to make a checkmark on
the table as you add each reagent. Cover each tube with Parafilm and invert twice to mix.
DO NOT MOUTH PIPET!
6. Thoroughly resuspend the mitochondrial suspension with a Pasteur pipet. Add the indicated volume
of mitochondrial suspension to each cuvet, recording the time added on Table 2. Remember to use
the boiled mitochondria prepared in step 3 (above) for tube 4. You will not be able to interpret
any of your results if you use the wrong suspension here. Check with your instructor if this is
not clear. Since you will not be able to add mitochondria to all 7 tubes simultaneously, you should
instead add mitochondria at definite intervals (for instance, every 30 sec). Be careful with your choice
of times so that you don't need to take two readings simultaneously. As soon as the mitochondrial
suspension has been added to a cuvet, cover the cuvet with Parafilm and invert to mix. Remove the
Parafilm and set the tube in the test tube rack provided.
7. After 5 minutes have elapsed from the time of addition of mitochondria to tube 1, set the
spectrophotometer to 100% T using blank 1 and take the 5 minute reading for tube 1. (Remember to
read absorbance when taking the measurement. Do not read from the percent transmittance scale. At
the appropriate time, adjust the spectrophotometer to 100% T with blank 2 and take the 5 minute
reading for tube 2. Then adjust the spectrophotometer to 100% T with blank 3 and take the 5 minute
readings for tubes 3 and 4. Enter the absorbance readings on Table 2 (p. 84).
80
MEASUREMENT OF SUCCINATE DEHYDROGENASE ACTIVITY
EXPERIMENTAL ASSAYS
The procedure above and Table 1 (below) describe how to set up an experiment to show that Vo, the
initial enzyme velocity, affects the enzyme velocity. For full credit, you must plan at least one additional
experiment that tests an idea about how different conditions affect succinate dehydrogenase activity.
How does changing the temperature affect the enzyme velocity?
What effect do different sugars involved in the citric acid cycle have upon SDH activity?
Is any succinate found in the mitochondria when they are isolated?
Is sodium azide required in the basic assay for DCIP to act as an electron acceptor?
What substances are rate-limiting components in the basic assay listed above?
You are not expected to test all of the ideas above, but need to include at least one-well formulated
experimental hypothesis in addition to the test described in the basic assay. Feel free to design other
experimental questions, but consult with your instructor well before the lab period if additional equipment
or reagents will be required. In addition to the materials listed for the basic assay, the following
equipment and reagents will be available. You should prepare a table similar to Table 1 listing the
components to be included in your additional assay before the lab period.
Styrofoam containers to be used to maintain constant temperature water baths.
0.2 M malonate
0.2 M fumarate
0.2 M citrate
Table 1. Assay Tubes for Succinate Dehydrogenase
Tube
Assay
Medium
Na Azide
(0.04 M)
DCIP
(5x10-4 M)
Succinate
(0.2 M)
Mitochondrial
Suspension
Blank 1
3.7 ml
0.5 ml
-------------
0.5 ml
0.3 ml
1
3.2 ml
0.5 ml
0.5 ml
0.5 ml
0.3 ml
Blank 2
3.1 ml
0.5 ml
-------------
0.5 ml
0.9 ml
2
2.6 ml
0.5 ml
0.5 ml
0.5 ml
0.9 ml
Blank 3
3.4 ml
0.5 ml
-------------
0.5 ml
0.6 ml
3
2.9 ml
0.5 ml
0.5 ml
0.5 ml
0.6 ml
4
2.9 ml
0.5 ml
0.5 ml
0.5 ml
0.6 ml
(boiled 5')
81
USING THE SPECTROPHOTOMETER
1. Rotate the power switch/zero control knob clockwise to turn on the spectrophotometer. A red light to
the left of the display should come on. Allow the machine to warm up at least 5 minutes before using.
2. Select the desired wavelength with the wavelength control.
3. Check to be sure that the sample compartment is empty and the cover is closed. Set the meter to 0%
transmittance using the zero control knob. Use the mirror behind the needle to avoid parallax errors
when reading the display.
4. Insert the reference blank. Use the transmittance/absorbance control to adjust the meter to 100%
transmittance.
Be sure to use the appropriate blank, as specified in the directions for your experiment. Some
experiments may require that you use different blanks, depending upon the sample that is
being measured. You will use three blanks in today's experiment, and will need to adjust to
100% transmittance for each one.
5. Insert the unknown sample. Read the absorbance from the display. Do not change any of the controls
before making this reading. Do not record the values on the percent transmittance scale.
Key operating features of the Spectronic® 20 Spectrophotometer. Figure courtesy of Milton Roy company.
82
CLEANUP
1. Wash the mortars and pestles immediately after use, dry them with paper towels, and return them to
the cold room. Be careful with these - they are very expensive.
2. Wash the centrifuge tubes and leave them inverted on the front bench in test tube racks to dry.
3. Wash the graduated cylinders you used and leave them inverted to dry on your bench.
4. Discard all solutions from the cuvets in the sink. Wash the cuvets and leave them inverted in the
plastic test tube rack on your bench.
5. Place used glass pipets in the pipet soaking container.
6. Put used plastic pipeter tips in the designated container.
7. Leave your bench area clean and neat.
REFERENCES
Bonner, Jr., W.D. (1967) A general method for the preparation of plant mitochondria. In Methods in
Enzymology, Vol. 10, Estabrook, R.W. and Pullman, M.E., eds. pp. 126-133. Academic Press, N.Y.
Lehninger, A.L. (1982) Principles of Biochemistry, Worth, NY
Stryer, L. (1988) Biochemistry, 3rd ed., W.H. Freeman & Co., NY
83
Laboratory 7
Spectrophotometric Assay of Mitochondrial SDH
Section ____________
Name: _____________________________________
(Friday AM/ Friday PM)
Lab Partners: _________________________________________________________________________
Table 2. Absorbance Readings (A600) At Each Time Interval
Tube
Time mitochondrial
suspension was added
1
0
2
30"
3
1'
4
1' 30"
5
min
10 min
15
min
20 min
25 min
30 min
Note: prepare an additional table to record measurements for your experimental assays.
Table 3 Total Change in Absorbance (ΔA600) At Each Time Interval
Tube
5 min
10 min
15 min
20 min
25 min
30 min
35 min
1
2
3
4
See next page for an explanation of ΔA600, and how to calculate these values.
Note: prepare an additional table to record measurements for your experimental assays.
84
35 min
LAB REPORT
Use the general guidelines provided at the beginning of the lab manual to write your report, which should
include a title, introduction, methods, results, and discussion. Specific points that should be covered in
your report are given below. Be sure to review the lab report guidelines, since points will be deducted for
not following the specified format. Your report must clearly describe the hypothesis and results for the
additional experiment you designed, and should include a clear discussion of these results. Remember that
Figures and Tables included in your report should be clearly labeled, and should include Figure or Table
legends (see lab report guidelines). You may use a computer graphing program to prepare the figures
required for this report; however, it is also acceptable to plot the graphs required by hand. In either case,
be sure to review the information in the introduction to the lab manual about presenting data in figures.
The experiment you design should be integrated into this report, rather than being presented as a separate
report.
Some points to help with lab report preparation:
1) In the methods section, values for centrifugal force should be reported in g force, rather than in
rpm values. This is because it is the g force that controls the sedimentation of cellular
components; a particular rpm value will give different g forces depending on the centrifuge rotor
(head) used.
2) The data tables on page 84 are an important part of the data collected and must be included in the
report. It is acceptable to include this page from your lab manual with values entered by hand.
Alternatively, you can retype the values to include in a table in your report. In either case, a
legend for each table should be included, as directed in the general guidelines for laboratory
reports.
3) Enter the total change in absorbance (ΔA600) for each tube in Table 3 (p. 84). The total change in
absorbance is the difference between the 5-minute reading for tube 4 and the A600 reading
for each tube at the specified time. For example, suppose the 5-minute reading for tube 4 was
0.73, and the readings for your experimental tube were 0.63 at 5 minutes and 0.42 at 10 minutes.
The ΔA600 for this tube at 5 minutes is then 0.73 - 0.63 = 0.10. The ΔA600 at 10 minutes is 0.73 0.42 = 0.31.
4) Use a sheet of linear graph paper (a master sheet to copy is provided at the beginning of the lab
manual) to plot the reaction rates for tubes 1-4. Plot the total change in absorbance (Y axis)
versus time (X axis). Include the origin as a point for all curves; that is, the 0 minute value for
ΔA600 is 0. Draw the best-fit curve for each plot with a ruler or French curve. Both axes of your
graph should be clearly labeled and indicate the units of measurement used. Provide a descriptive
title for your graph, and draw the four plots in a way that makes them easy to distinguish. For
example, you could use different symbols for different conditions (e.g. ■, ▲, or ●), and use
dashed, solid, or colored lines to distinguish the plots. To make it easier to compare the reaction
rate for the different volumes of mitochondria, the reaction rates for tubes 1-4 should all be
plotted on a single graph. A figure legend should also be provided for each graph, briefly
summarizing the data presented and explaining the differences between the reactions presented in
the four tubes. Labels should be informative - for example, labeling a line as "growth after
treatment with estrogen" is more helpful than labeling the same line "Tube 3". You may plot the
graph using a computer if you choose - if you do so, remember that the plot is expected to go
through the origin, and that your axes and lines need to be clearly identified. You may need to
alter the program defaults to allow this.
85
5) Use a second piece of graph paper to plot the initial velocity (Y axis) versus enzyme
concentration (X axis) for tubes 1-3. The intial velocity (V0) is the rate at which ΔA600 changes
per unit time at the beginning of the reaction. Since the velocity may change during the reaction,
one way to determine V0 is to take the total change in absorbance after 5 minutes (from the graph
made as described above) and divide by 5 to obtain the ΔA600/min. However, you will need to
inspect your graph to determine if a longer time period would be better for determining V0. For
the different enzyme concentrations, simply use the three volumes of mitochondrial suspension
added: 0.3 ml, 0.6 ml, and 0.9 ml mitochondria/reaction. The origin should be used as a fourth
point. Draw the best-fit straight line for the four points. The theoretical prediction (from
Michaelis-Menten kinetics) is a straight line; drawing a straight line on the graph will allow you
to see how much the observed data differs from the expected. Again, provide informative labels
and a figure legend.
6) Remember that your results section must include a narrative to provide context for the data
presented in the figures. See the guidelines to lab reports at the beginning of the lab manual for
more information.
7) After presenting your results, the following questions should be addressed as part of your
conclusions. The conclusions should be written in complete sentences and organized in
paragraphs. They should be written in an organized fashion so that ideas flow logically.
Remember that the conclusions should focus on the experimental results you observed, rather
than broad generalizations that don’t relate specifically to your experiments.
a) What volume of mitochondrial suspension gave the highest initial velocity? Comment on the
relationship of Vo and enzyme concentration.
b) You may have observed that the reaction rate was much lower at the end of the reaction time than
at the beginning for some of the reactions. If so, this suggests that one of the reaction components
was no longer present at a high enough concentration for the reaction to proceed at a maximal
rate. What compound included in the reaction mixture do you think was the rate limiting
component? Present evidence that supports your hypothesis, or suggest an additional experiment
that could be used to support the hypothesis you made.
c) What effect did boiling the mitochondria have upon SDH activity? Why was this effect observed?
d) Why could the 5 minute absorbance reading for tube 4 be assumed to be the same as the 0 minute
reading for all tubes for calculation of the ΔA600?
86
LAB CRITIQUE SHEET
Laboratory 7 Spectrophotometric Assay of Succinate Dehydrogenase
Clarity of written introduction/background
1 2
Unclear
Length of written introduction/background
Too short
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1
Clarity of procedures
1
Rate this lab relative to
other labs at NMU
3
5
4 5
Very
Clear
2 3 4
Too long
5
1 2
Unclear
3
4 5
Very
Clear
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Low
3
4
5
High
Relevance to lecture material
Rate this lab relative to
other BI 313 labs
4 5
Very
Clear
2 3 4
Too long
1 2
Unclear
Length of in-lab introduction/background
Too short
3
1
Poor
2
3
4
5
Excellent
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Poor
2
3
4
5
Excellent
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88
LABORATORY 8
CELL MOTILITY AND THE CYTOSKELETON
One of the distinguishing features of eukaryotic cells is the presence of different types of cytoskeletal
proteins. These proteins provide for specific cell shapes, can keep organelles in specific locations in the
cell, and can also allow some cells to move. Although one usually thinks of animal cells moving, plants
are also capable of movement. Cytoplasmic streaming within a plant cell can carry organelles around the
cytoplasm; this streaming is mediated by cytoskeletal proteins. Movement of chromosomes during
mitosis in plant cells uses the same cytoskeletal proteins that are used during mitotic movements in
animal cells. Some unicellular plants and many lower plant sperm cells are flagellated. In addition to
these movements, which involve cytoskeletal proteins homologous to those found in animal cells, some
plant movements involve changes in turgor pressure. For example, guard cells around stomata change
shape to control the size of the leaf openings; some flowers open and close during the day; and sensitive
plants respond to touch by closing their leaves; plants can grow toward a source of light.
There are several different motor proteins that mediate cell movement. Actin and myosin interact in
muscle contraction and are also involved in amoeboid movement. Tubulin and other proteins in the
axoneme are responsible for the movement of cilia and flagella, and tubulin and a motor protein(s) still to
be characterized are needed for chromosomal movements in mitosis. Kinesin and cytoplasmic dynein
move vesicles along microtubules in axons.
In today's laboratory you will observe cell motility in several different unicellular organisms. These
organisms are relatively large and robust enough to withstand a reasonable amount of experimental abuse.
We will observe amoeboid movement in the amoeba Chaos chaos, ciliary motion in Paramecium
caudatum, and flagellar motion in the alga Carteria crucifera.
89
AMOEBOID MOTION
Amoeboid motion is observed in many protozoa, in fungi, in animals, and even in the sperm of some
plants. Amoeboid motion is a common feature in animal embryonic development and is retained in a few
adult cells, such as white blood cells. Other adult cells, like fibroblasts, may migrate into the region of a
wound to aid healing. Normally non-motile cells may become capable of amoeboid movement if they are
transformed into cancer cells.
Amoeboid cells extend cytoplasmic lobes called pseuodopods in the direction of cell movement. Careful
examination of the pseudopod shows that the region around the cortex of the cell is in a semisolid, or
gelled state. Liquid cytoplasm can be seen streaming in the direction of movement through the
pseudopod.
Even though amoeboid movement is easy to observe with a light microscope, the mechanism is not well
understood. Several points of evidence implicate actin and myosin in this type of movement. First, both
actin and myosin can be extracted from cells capable of amoeboid movement. Second, electron
micrographs reveal thin filaments in the region of the cell cortex. Immunofluorescence studies show that
both actin and myosin are found in this region. Third, treating cells with drugs like cytochalasin or
phalloidin that interact with actin filaments, will stop amoeboid motion. D.L. Taylor and his coworkers
(for example, see Condeelis and Taylor, 1977) have proposed that contraction of the cortical actin gel in
the rear and midregions of an amoeba forces the liquid cytoplasm through the pseudopod towards the tip.
This contraction is thought to mediated by actin-myosin interactions. Assembly and disassembly of actin
filaments in the cortical gel also occur during amoeboid movement, with disassembly occurring
principally at the rear end of the cell.
CILIARY AND FLAGELLAR MOVEMENT
The axoneme (see Fig. 16-51, MBOC) is found in the core of both cilia and flagella. Nine doublet
microtubules form a circle around a pair of single microtubules. Dynein, the motor protein in cilia and
flagella, extends from each of the doublet microtubules to make contact with an adjacent doublet. When
cilia and flagella bend, the dynein arms cause the adjacent microtubule doublets to slide relative to one
another. Other accessory proteins in the axoneme regulate the direction of bending and prevent the
axoneme from sliding apart. (Remember that cilia and flagella are plural nouns; the singular names of
these structures are cilium and flagellum.)
Axonemal proteins are not capable of self-assembly into their organized structure, but require a structure
to serve as a template. This nucleating structure in the cell is the centriole; at the base of cilia and flagella,
the centriole is traditionally called the basal body. A separate basal body is found at the base of each
cilium or flagellum in the cell. The organization of the basal body is similar to the axoneme, but the outer
microtubules are found as triplets rather than doublets (see Fig. 16-45, MBOC).
Cilia are relatively short (5-10 µm long) and are abundant on the surface of a cell, while flagella are long
(up to 150 µm) and are usually found singly or in pairs, although sometimes a few flagella will be found
on a cell. Cilia typically bend from the base to make an oarlike power stroke, and then return to the
original position in a recovery stroke, while flagella typically bend in a sinusoidal wave. Both beat
patterns can be observed in the flagella of the alga Chlamydomonas, where the two flagella beat in a
ciliary pattern to move the cell forward and change to the flagellar pattern to move the cell in reverse.
90
Observations of ciliary movement in Paramecium
We will use the protozoan Paramecium caudatum to study ciliary movement. This organism uses cilia
both for locomotion and to move food into the buccal cavity. The rapidly swimming Paramecia will be
slowed down by adding yeast to a wet mount; as the Paramecia gather around air bubbles and clusters of
yeast cells they will slow down and will be easier to observe. The yeast cells will have been stained with
the dye Congo red to provide contrast. A diagram showing the major features of Paramecium will be
provided in lab.
Observations of flagellar movement in Carteria
Carteria is a motile alga with four flagella at one end of the cell. You will make two wet mounts of
Carteria — one in the culture medium in which the alga was grown, and one with a drop of Protoslo
(Carolina Biological) added. Protoslo is a viscous liquid that will slow down the swimming cells to make
them easier to observe; however, it may also affect the movement of the flagella. A diagram showing the
general appearance of Carteria will be provided.
Regeneration Of Carteria Flagella
The flagella on Carteria can be removed experimentally. The cells will then regenerate flagella if they are
transferred to fresh liquid medium. Although we will not experiment with flagellar regeneration in this
laboratory, you might consider flagellar regneration for your research project. Rosenbaum et al. (1969)
describe flagellar regeneration experiments with Chlamydomonas, another species of flagellated algae.
References
Condeelis, J. S., and Taylor, D. L. (1977). The contractile basis of amoeboid movement. V. The control of
gelation, solation, and contraction in extracts from Dictyostelium discoideum. J. Cell Biol. 74, 901-927.
Rosenbaum, J. L., Moulder, L. E., and Ringo, D. L. (1969). Flageller elongation and shortening in
Chlamydomonas: The use of cycloheximide and colchicine to study the synthesis and assembly of
flagellar proteins. J. Cell Biol. 41, 600-619.
91
PROCEDURES
MICROSCOPIC EXAMINATION OF Chaos chaos
Students should work in groups of three to prepare slides of amoeba for observation, as directed below.
1. Use a dissecting microscope to locate the amoebae on the bottom of the culture dish. Bring the tip of
a Pasteur pipet next to an amoeba, and carefully draw it into the tip of the pipet. Only 2-3 cm of liquid
should be drawn into the pipet. Expel the liquid onto a clean microscope slide. Chaos chaos is large
enough that you should be able to see if the transfer was successful with the naked eye. Use the
dissecting microscope to confirm that you do indeed have an amoeba on the slide.
2. To avoid crushing the amoeba, you need to prepare an elevated culture chamber for observation.
Spread a thin film of petroleum jelly between your thumb and forefinger. Gently scrape all four sides
of a cover slip against this film to make a thin rim of petroleum jelly. Carefully lower the cover slip
on top of the amoeba prepared in step 1. Avoid making air bubbles, since these will interfere with
your observations.
3. Use phase contrast optics and 100X final magnification to observe the amoeba and watch its
movement. It may take a few minutes after the transfer before the amoeba begins moving. Note the
direction of movement and extension of pseudopods.
4. Observe the cytoplasmic streaming in different parts of a pseudopod using 100X, 200X, and 400X
final magnification. Remember that you need to rotate the condenser to the appropriate position as
you increase magnification when using phase contrast. Do not try to use the oil immersion lens for
observation, since the culture chamber you prepared is too thick to fit under this lens. Note any
differences observed in different portions of the pseudopod.
5. Draw a sketch of an amoeba in the space provided on page 69. Use your ocular micrometer to
measure the length of the amoeba, and indicate its size (in µm) and the final magnification used for
observations on your drawing.
6. Compare the amount of detail that you can see using phase- contrast optics to what you can observe
with bright-field optics.
92
MICROSCOPIC OBSERVATION OF Paramecium
1. Place a drop of the Paramecium culture in the center of a clean slide. Dip a toothpick into the Congo
red-stained yeast suspension and then dip this toothpick into the Paramecium culture on your slide.
Only a small amount of yeast is needed — if the drop turns red rather than pink, you added too much.
Put a thin film of petroleum jelly on all four edges of a coverslip, and place the coverslip on top of
your culture.
2. Use 100X final magnification to locate an area of the slide with Paramecia. They will be much larger
than the yeast cells and will be swimming quite rapidly initially. As the Paramecia begin feeding on
the yeast, they will slow down. Find a slow-moving specimen and observe the movement of the cilia
as it swims and as it feeds on yeast. (Don't be confused by other, smaller, ciliated protozoa that may
be present - if you are not sure, check your text for the expected size for a Paramecium).
Note: Although you may find phase-contrast optics useful for these observations, you should not
use the green filter, since that would make it impossible to distinguish the color of the Congo-red
stained yeast cells.
3. The yeast cells are collected in a food vacuole at the anterior end of the animal. The dye Congo red,
which was used to stain the yeast, is a pH indicator. At pH 5 and above, this dye is bright red,
between pH 3 and 5 it is purple, and below pH 3 it is blue.
4. Draw a sketch of a Paramecium in the space provided on page 94. Use your ocular micrometer to
measure the length of the organism and indicate its size (in µm) on your drawing. Label the drawing
with the organism observed and indicate the final magnification used for observation.
MICROSCOPIC OBSERVATION OF Carteria
1. Use a Pasteur pipet to remove a small drop of the Carteria culture from the top of the culture jar,
where the majority of motile cells are located. Place the drop in the center of a clean microscope slide
and cover with a coverslip (no petroleum jelly is needed for this slide). Observe the normal swimming
behavior of Carteria using 100X final magnification.
2. Add a drop of Carteria to a fresh slide and mix in a small drop of Protoslo. Mix with a toothpick and
add a coverslip.
3. Examine using 100X, 200X, and 400X final magnification. If you adjust the illumination carefully,
you should be able to see the flagella. Experiment with both bright-field and phase-contrast to see
which gives a better image of the flagella.
93
Drawings
Organism observed: ___________________
Final magnification: ________
Organism observed: ___________________
Final magnification: ________
94
LAB CRITIQUE SHEET
Laboratory 8 Cell Motility and the Cytoskeleton
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Rate this lab relative to
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Relevance to lecture material
Rate this lab relative to
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4 5
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2 3 4 5
Too long
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3
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96
LABORATORY 9
ACTIN-BASED CELL MOTILITY: MYOFIBRIL CONTRACTION
Background Reading:
Molecular Biology of the Cell, pp. 847-858
INTRODUCTION
One of the best understood contractile systems is the myofibril. The highly organized actin and myosin
filaments that are present in the myofibril have allowed investigators to study the cellular basis of muscle
contraction. In this laboratory, you will use phase contrast microscopy to observe myofibril contraction,
and will add different solutions to test the biochemical requirements for such contraction.
The myofibrils you will use for this experiment have been isolated from rabbit psoas muscle. This tissue
was selected because the myofibrils are long and straight, and are relatively free of connective tissue. The
tissue was treated with glycerin, which disrupts the membranes of the muscle cells, allowing ATP and
inorganic ions to leach out of the cells. Since the cell membranes have been permeabilized, you will be
able to add different solutions to test their effect upon muscle contraction. The glycerination process does
not disrupt the highly stable array of actin and myosin in the myofibril, so contraction of the cells will still
occur.
EXPERIMENTAL PROCEDURES
Students should work in pairs for this experiment. A supply of clean microscope slides, cover glasses, and
Pasteur pipets will be available on the front bench. If the microscope slides are not brand new, wash them
using detergent from the small jar in the top drawer of your bench, rinse carefully, and dry with tissues.
Each row of students will be supplied with an ice bucket containing different test solutions. You will also
need a compound microscope, adjusted properly for phase contrast optics. (See Laboratory 1 if you need
a review on using phase contrast.) Each bench should also have a supply of filter paper strips to be used
for the perfusion technique.
1. Obtain an aliquot of a myofibril suspension from the ice bucket on the front bench. A single 1-ml
aliquot should be sufficient for all of the contraction experiments. Allow the preparation to warm to
room temperature before beginning the contraction experiments. The myofibril preparation can be
kept at room temperature during the 3-hour laboratory period, but was stored on ice for long term
stability.
2. Agitate the myofibril suspension gently by hand. Using a Pasteur pipet attached to a rubber bulb,
transfer a small drop to the center of a microscope slide. Apply a cover glass to the preparation. The
solution should flow freely to the edges of the slide, but the cover slip should not slide from side to
side. Each student should prepare a myofibril slide, so independent drawings can be made for part 4.
97
3. Focus on the myofibrils, beginning with the 100X magnification, and then moving to 200X and
400X. You should use phase-contrast optics, remembering to change the condenser and to properly
align the phase rings for each magnification. Apply a small drop of immersion oil to the center of the
field of view (look for the illuminated circle on the slide). Carefully swing the 100X objective into
place. There should be a continuous film of oil between the cover slip and the objective lens. Use the
fine focus knob to bring the myofibrils into sharp focus. You may need to adjust the illumination for
optimal contrast.
Problems to watch for:
1)
Be careful that the objective lens does not crash into the slide or cover slip.
2)
Too much liquid can cause a fuzzy image. If you suspect this is the case, try wicking
some of the extra liquid away by touching a strip of filter paper to the edge of the cover
slip.
3)
If oil seeps under the cover slip, poor contrast will result. Focus on myofibrils near the
center of the slide, so you won't have to apply immersion oil near the edge.
4. Make a sketch of the myofibrils as observed with 1000X final magnification and phase contrast
optics. Under phase contrast, I-bands appear light and A-bands appear dark. You should also be able
to observe the Z-discs as dark lines in the center of each I-band. Finally, clearly indicate the extent of
a single sarcomere in your drawing. Your drawing should be clearly labeled, and based on your own
observations. Note that you will not be able to see all the details shown in the electron micrographs of
sarcomeres shown in your text.
5. Measure the length of four sarcomeres from different myofibrils, and record the values in ocular
division units. Taking the average from multiple measurements provides a more accurate value than a
single measurement. After calculating the average, convert this value to micrometers, using the
conversion value from lab 1. These measurements will tell you the average length of a sarcomere in
the relaxed state.
6. Each pair of students should then prepare a fresh wet mount of the myofibril suspension, and focus
upon the myofibrils using 400X magnification and phase-contrast optics. Using a Pasteur pipet
attached to a rubber bulb, transfer five drops of Perfusion Solution I (ATP/Mg2+/Ca2+, see Table 1) to
one edge of the cover slip. Apply a strip of filter paper to the opposite side of the cover slip. The filter
paper will act as a wick, removing the old solution and drawing in Perfusion Solution I. One partner
should observe the myofibrils through the microscope while the other does the perfusion, refocusing
with the fine focus as necessary. If the perfusion is done properly, you will see myofibrils moving
across the field of vision. After the myofibrils begin moving, wait 15-20 seconds, and then direct your
partner to remove the paper wick. Make complete written descriptions of the appearance of the
myofibrils after treatment with Perfusion Solution I, providing a qualitative description of the changes
observed as this solution was added. You may need to wick excess liquid from under the cover slip to
provide a sharp image.
7. Repeat the procedure in step 6 for the other six perfusion solutions (II through VII) listed in Table 1.
For each perfusion solution. be sure to do the following:
a. Make thorough written descriptions of the treated myofibrils and their responses and
changes, if any, to the perfusion solution.
b. Observe the myofibrils as perfusion takes place. Complete contraction, when it occurs,
takes place rapidly. If you have enough myofibril suspension, you may wish to repeat the
more interesting perfusions so each partner can watch contraction while it occurs.
98
LAB REPORT
A formal report is not required for this laboratory. Instead, you should turn in:
A A table showing the sarcomere length measurements prior to contraction.
A A labeled drawing of myofibrils in the relaxed state.
A A qualitative description of the changes observed after each perfusion solution was added, and a
brief explanation of the effects observed.
b
Use the information from your observations to answer the following question. The physiological
substrate for many enzymes that hydrolyze ATP is a complex of Mg2+ and ATP, often referred to
as MgATP. Do your observations indicate that the myosin ATPase requires MgATP, or is ATP
alone sufficient?
CLEANUP
1. Used cover slips can be discarded in the broken glass container at the front of the room.
2. Microscope slides should be washed and returned to the front bench.
3. Pasteur pipets should be placed in the soaking container on the front bench.
4. Be sure all perfusion solutions are returned to the front bench.
5. Any leftover myofibril suspension from the aliquot you kept at room temperature can be discarded.
6. Return the microscope condenser to the "0" position, put the low-power (10X) objective in place, and
return your microscope to the appropriately numbered space in the cabinet.
7. Leave your bench area clean and neat.
REFERENCES
Franzini-Armstrong. C. and Peachey, L.D. (1981) Striated muscle: contractile and control mechanisms. J.
Cell Biol. 91 (3, Part 2):166s-186s.
Gasque, C.E. (1989) A Manual of Laboratory Experiences in Cell Biology Wm. C. Brown Publishers,
Dubuque.
Pollard, T.D. (1981) Cytoplasmic contractile proteins. J. Cell Biol. 91 (3, Part 2):156s-165s.
99
Table 1
Perfusion solutions used for the study of myofibril contraction.
Perfusion solution
Designation of Perfusion Solution
Composition of Test Solution
I
ATP/Mg2+/Ca2+
100 mM KCl, 0.5 mM ATP; 1 mM
MgCl2; 4.1 mM CaCl2; 10 mM
histidine-HCl, pH 7.0
II
No ATP/Mg2+/Ca2+
100 mM KCl; 1 mM MgCl2; 4.1
mM CaCl2; 4 mM EGTA; 10 mM
histidine-HCl, pH 7.0
III
ATP/No Mg2+/No Ca2+
100 mM KCl, 0.5 mM ATP; 4 mM
EGTA; 10 mM histidine-HCl, pH
7.0
IV
ATP/Mg2+/No Ca2+
100 mM KCl, 0.5 mM ATP; 1 mM
MgCl2; 4 mM EGTA; 10 mM
histidine-HCl, pH 7.0
V
ATP/No Mg2+/Ca2+
100 mM KCl, 0.5 mM ATP; 4.1
mM CaCl2; 10 mM histidine-HCl,
pH 7.0
VI
MES (Myosin Extraction Solution);
dissociates myosin, but not actin, from
the myofibrils.
0.6 M KCl; 1 mM MgCl2; 0.01 M
sodium pyrophosphate, pH 6.5
VII
AES (Actin Extraction Solution);
dissociates both myosin and actin from
the myofibrils.
0.6 M KSCN, 6.67 mM potassium
phosphate, pH 7.0
100
Laboratory 9
Actin-based cell motility: myofibril contraction
Name ____________________________________
Section ______________
Friday AM/ Friday PM
Table 2. Sarcomere Length Measurements
Sarcomere
length
Sarcomere
length
Sarcomere
length
Sarcomere
length
101
Average
sarcomere
length
(divisions)
Average
sarcomere
length (µm)
Skeletal muscle cell (muscle fiber) from a vertebrate.
Fig 16-82, Molecular Biology of the Cell Alberts, Bray, Lewis, Raff, Roberts, and Watson, 1994.
Garland Publishing, N.Y.
Diagram of myosin and actin filaments of a sarcomere.
Fig. 16-89, Molecular Biology of the Cell Alberts, Bray, Lewis, Raff, Roberts, and Watson, 1994.
Garland Publishing, N.Y.
102
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LABORATORY 10
MITOSIS AND THE CELL CYCLE
The cell cycle can be divided into M phase when the cell is actively dividing and the chromosomes are
condensed, and interphase, the period between cell divisions. M phase includes two overlapping sets of
events, mitosis and cytokinesis. Both mitosis and interphase can be further subdivided. Interphase
consists of the G1 phase, the period after cell division has been completed and before DNA replication
begins; S phase, during which DNA is replicated and two sister chromatids are assembled; and G2
phase, the interval between the completion of DNA replication and the initiation of mitosis. (See Fig.
17-3, MBOC).
Mitosis is divided into several stages which are illustrated in Panel 18-1 (pp. 916-917) of Molecular
Biology of the Cell. The transition between G2 phase and prophase is not sharply defined. During
prophase the chromosomes condense, cytoplasmic microtubules disassemble, and the mitotic spindle
begins to form. At the end of prophase condensed chromosomes will be visible in an intact nucleus. The
initiation of prometaphase is marked by the breakdown of the nuclear envelope. The mitotic spindle
assembles and moves the chromosomes from random locations towards the middle of the cell. In a cell in
metaphase, the chromosomes are aligned along the metaphase plate in the center of the cell. Anaphase
begins as the sister chromatids separate and are drawn apart to opposite spindle poles. During telophase,
the condensed chromosomes begin to decondense and the nuclear envelope reforms.
Several kinds of experiments that have been done to study the different parts of the cell cycle are
described in Molecular Biology of the Cell, pp. 864-867. The easiest parts of the cell cycle to observe are
the different stages of mitosis, since dramatic events involving chromosome condensation and movement
are easily observed with the light microscope. In this week's lab you will stain root tips from the broad
bean, Vicia faba, with a cytochemical stain that is specific for DNA. After scoring cells that are in the
different stages of mitosis, you will be able to calculate the time it takes the cells to complete each stage.
Vicia faba has been chosen for this experiment because this organism has a small number of relatively
large chromosomes. This makes it easier to spread the chromosomes and observe their morphology on a
microscope slide. In V. faba there are twelve chromosomes in the diploid state: five pairs of acrocentric
chromosomes and one pair of telocentric chromosomes. Chromosomes are called either acrocentric,
metacentric, or telocentric, depending on the position of their centromere and the relative lengths of the
chromosomal arms. Figure 1 illustrates these three general types of chromosomes.
105
Figure 1
Chromosome types
The centromere of each chromosome is readily recognized by the constriction that forms where the two
sister chromatids are attached in prophase or metaphase cells. In some chromosomes, another
constriction, called the secondary constriction, is visible. The secondary constriction is a very thin,
lightly stained region that may appear as a gap within the chromosome arm. This region is the site of the
nucleolar organizer, the DNA from which ribosomal RNA is transcribed. In V. faba, one pair of
chromosomes has a secondary constriction.
FEULGEN REACTION
Cell biologists often wish to stain specific kinds of macromolecules within cells so they can tell exactly
where those molecules are located within the cell. This procedure is known as cytochemistry. We will
use a cytochemical reaction known as the Feulgen reaction, introduced by R. Feulgen and H. Rossenbeck
in 1924, to stain the chromosomes in dividing root tips. Although we will simply be using this reaction to
make the chromosomes easier to see, the Feulgen reaction can also be used to quantify the amount of
DNA in individual cells using a procedure called microspectrophotometry.
The root tips that you will stain have been fixed in 1:3 acetic acid:ethanol and stored in 70% ethanol.
Fixation makes cells permeable to the chemicals used for staining and crosslinks macromolecules within
the cell so they are fixed in position. Fixation also inactivates enzymes that might otherwise degrade the
tissue to be examined. See p. 45 for more information.
In the first part of the Feulgen reaction, the root tips will be treated with hot HCl. This hydrolyses the
DNA molecules in the chromosomes, removing purines and leaving an exposed aldehyde group attached
to the depurinated deoxyribose sugars (see Fig. 2). The root tips are then treated with Schiff's reagent, a
clear liquid prepared by treating a red dye, basic fuchsin, with sulfurous acid. The fuchsin reacts with free
aldehydes to form a reddish purple product. RNA is not stained in this reaction since all ribonucleic acid
in the cell is removed during the HCl hydrolysis step.
106
The specificity of the Feulgen reaction comes from the HCl hydrolysis, which removes purines from the
DNA backbone, exposing free aldehydes. As you saw earlier in the semester, Schiff's reagent can also be
used to stain cellular carbohydrates if a different chemical reaction (using periodic acid) is used to oxidize
sugars to create free aldehydes.
SQUASH PREPARATIONS
The tissues that you will be staining in this lab are V. faba root tips. At the very tip of the root is a region
called the apical meristem, where actively dividing cells are found. If the entire root tip were placed under
the microscope, it would be too thick to see individual cells. Although sectioning a tissue is one common
way to get a thin enough slice to observe with a microscope, in a section only part of the cell is present. If
we were to section the root tips, many cells cut by the section would not have the plane of section in the
nucleus. Even for those cells cut within the nucleus, it would be impossible to see all the chromosomes in
the cell. By squashing the cells flat on the slide, you can spread all the chromosomes from a single cell
out in a thin layer. Squashing also disperses the chromosomes on the slide to make it easier to count them
and observe their morphology. This procedure is frequently used to examine chromosomes from both
plant and animal tissue.
MEASURING THE DURATION OF MITOSIS
If a population of cells is dividing asynchronously, the percentage of cells that are in mitosis at a
particular instant is the same as the percentage of time required for completion of mitosis. The percentage
of cells in mitosis is called the mitotic index. Therefore, the length of time required to complete mitosis is
approximately equal to the mitotic index multiplied by the time required for a complete cell cycle. The
same principal can be applied to calculate the duration of each stage of mitosis. The values you obtain
will not be completely accurate, since not all cells in the root tip are mitotically active. However, the
values obtained should give a reasonable estimate of the time required to complete the different stages of
mitosis.
107
Figure 2. Feulgen Reaction. From Bregman (1990), Laboratory Investigations in Cell and Molecular
Biology, J. Wiley & Sons, N.Y.
108
PROCEDURES
WARNING: Although Schiff's reagent is colorless, or nearly so, it will stain
fingertips, clothing, notebooks, and anything else it contacts a brilliant purple
color.
Each row of students should work together to carry out the Feulgen reaction on the root tips. Once the tips
have been stained, students should work individually to squash the root tips and score the mitotic cells.
Note: whenever the root tips are handled, grasp the broad end of the root tip with forceps. The
mitotically active cells are at the narrow end, which should not be crushed.
FEULGEN REACTION
1. Collect one root tip for each student in the row in a beaker of distilled water. Collect two extra tips as
spares in case some are lost during the staining procedure. For example, for six students, stain eight
root tips.
2. Rinse the fixed root tips with several changes of distilled water. Carefully pour off the distilled water
while retaining the tips with forceps.
3. Grasp the broad end of each root tip with forceps and transfer the root tip to a beaker of 1 N HCl in a
60º C water bath. Rinse the forceps with water immediately.
4. After 10 minutes, remove the beaker from the 60º C water bath. Carefully add distilled water to dilute
and cool the acid.
5. Carefully pour off the diluted HCl and rinse the root tips with distilled water.
6. Use forceps to transfer the root tips to a small beaker containing Schiff's reagent. Rinse the forceps
with water. Cover the beaker with a coffee can to protect from light and keep in the dark for 30
minutes.
7. Use forceps to transfer the root tips to a beaker containing distilled water. Rinse the forceps in water
after the transfer. The intensely stained region near the tip of the root contains the mitotically active
cells.
109
SQUASH PREPARATION
1. Carefully clean a microscope slide, including a final wash with ethanol to make sure all grease is
removed.
2. Place a drop of 45% acetic acid in the center of the slide.
3. Grasp the broad end of a root tip with forceps and transfer the root tip to the drop of acetic acid.
4. Use a single-edge razor blade to cut off all but the terminal 1.5 mm of the root tip, which should be
intensely stained. Discard the unstained portion, and allow the stained portion of the root to remain in
the acetic acid for about 1 minute.
5. Gently tap the root tissue with a glass rod until you have a homogenous suspension, with no large
clumps of tissue. Gently lower a coverslip on the preparation.
6. Place the slide on a piece of paper towel. Tear off a piece of paper towel larger than the slide and
carefully place it on top of the coverslip. Hold this piece of towel in place with one hand, and with the
other hand press very gently with an index finger to press out excess acetic acid. It is very important
that the coverslip stay in position, rather than moving laterally. Lateral movement will cause the cells
to fold on top of one another, and make it very difficult to see flattened cells.
7. Tear off a fresh piece of paper towel and put it on top of the slide. Place your thumb in the center of
the cover slip and use a rolling motion to press liquid to the side of the cover slip. Use several rounds
of pressure, beginning with gentle pressure, and increasing to fairly firm pressure with the thumb. Be
careful that the coverslip does not slip from side to side.
8. After applying pressure with your thumb, carry out a final round of squashing with the eraser of a
pencil, using a new pencil to provide a flat surface. Again, be careful that the coverslip does not move
laterally.
9. Seal the edges of the coverslip with vaseline.
After you have stained your slide, use 100X magnification to locate a region of mitotically active cells
near the tip of the root. Examine these cells with the 400X magnification; for some questions you may
need to use the 1000X magnification (remember to use immersion oil). Answer the questions below and
count the number of cells that are in the different stages of mitosis, as directed. A formal report is not
required.
110
Laboratory 10 Mitosis and the Cell Cycle
Section ____________
Name _____________________________________
(Friday AM/ Friday PM)
1. Find several cells in interphase. Is the nucleolus visible? How does the staining intensity of the
nucleolus compare to the rest of the nucleus? Considering that the Feulgen reaction is specific for
DNA, what do the staining characteristics of the nucleolus suggest about its chemical composition?
You should be able to integrate your observations and your knowledge of the nucleolus and its
function to respond to this question.
2. Locate a cell that is in late prophase or early metaphase where the chromosomes are clearly spread
apart. How many chromosomes can you count in this cell? Draw a diagram of a chromosome showing the
appearance of the secondary constriction. [Do the best job you can to count the chromosomes, which are
likely to be fairly tangled. The number of human chromosomes was incorrectly reported as 48 for many
years because of this problem. Since Vicia faba has many fewer chromosomes, your task is easier.]
111
Laboratory 10 Mitosis and the Cell Cycle
Section ____________
Name _____________________________________
(Friday AM/ Friday PM)
3. Use the space below to draw a diagram of a cell showing well-spread chromosomes. Remember to
include the species observed and the final magnification used for observation in your drawing.
4. Score 200 cells for their mitotic stage, entering the data in the table below. The top line is provided to
enter data with hatch marks; these can be converted to numbers when you have collected data from
200 cells. Each student should score 200 cells from her/his own slide.
Interphase
Prophase
Metaphase
Anaphase
Telophase
Since some of the mitotic stages will have relatively few cells, it will be more accurate to collect data
from the entire class to calculate the duration of the different stages. Use the table on page 114 for the
class data.
112
Laboratory 10 Mitosis and the Cell Cycle
Section ____________
Name _____________________________________
(Friday AM/ Friday PM)
5. Calculate the mitotic index for the class, using the data from the class table.
____
Total number of cells scored
(interphase + all mitotic stages)
____
Total number of cells in mitosis
(prophase + metaphase + anaphase + telophase)
Mitotic Index
Number of cells in mitosis
Total number of cells
Mitotic index = _________
6. Use the value of the mitotic index calculated for question 5, and the known duration of the cell cycle
in Vicia faba (19 hr) to estimate the duration of mitosis.
Duration of mitosis __________ hr
7. Repeat the calculations above to calculate the duration of the different stages of mitosis. Show your
work on a separate piece of paper.
Duration of prophase:
______ hr
Duration of metaphase:
______ hr
Duration of anaphase:
______ hr
Duration of telophase:
______ hr
113
Laboratory 10 Mitosis and the Cell Cycle
Section ____________
Name _____________________________________
Interphase
Prophase
(Friday AM/ Friday PM)
Metaphase
Anaphase
Telophase
Total
Note: record total numbers for the class for each mitotic stage in the bottom row of the table.
114
LAB CRITIQUE SHEET
Laboratory 10 Mitosis and the cell cycle
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