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Laboratory Manual for Cell Biology

Biology 350 Department of Biological Sciences Salisbury University

By

E. Eugene Williams Associate Professor Department of Biological Sciences Salisbury University

Contents

Chapter 1. Guidelines for Writing a Scientific Paper. With emphasis on the preparation of Cell Biology Laboratory reports. Chapter 2. Pipetting and Creation of a Standard Curve. Chapter 3. Cell Fractionation: Isolation of Mitochondria from Cauliflower. Chapter 4. Cell Fractionation: Assay of Mitochondrial Enzyme Activity. Chapter 5. Cell Fractionation: Enzyme specific activity. Chapter 6. Enzyme Kinetics. Chapter 7. Protein Fractionation: Purification of IgG from Human Serum. Chapter 8. Protein Fractionation: Sodium Dodecyl Sulfate ­ Polyacrylamide Gel Electrophoresis. Chapter 9. Protein Fractionation: Western Blotting Chapter 10. Cell Culture: Determining the Density of Cell Cultures. Chapter 11. Cell Culture: Primary Culture of Chick Embryo Fibroblasts. Chapter 12. Cell Culture: Secondary Cultures of Chick Embryo Fibroblasts. Chapter 13. Cell Culture: Tumor Cell Cultures. Appendix 1. Reagents and Materials. Appendix 2. Instructions to Authors. Appendix 3. Manuscript ­ "Before". Appendix 4. Manuscript ­ "After". 2 13 19 25 32 36 42 50 55 58 65 71 74 78 85 88 88

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Chapter 1.

Guidelines for Writing a Scientific Paper.

With emphasis on the preparation of Cell Biology Laboratory reports.

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Objective This Chapter provides both general and specific guidelines for preparing a manuscript for publication in the scientific literature. These are guidelines and the format you will follow to prepare your Laboratory Assignments for this course. Follow these carefully and refer to them often while preparing reports. Use them later in your career to write scientific manuscripts. Introduction In Cell Biology (Biol 350), we are going to give you what is probably your first experience writing a paper in the style used by scientists. This is neither an English nor a Chemistry course, so much of this will be unfamiliar to you. Learning how to communicate effectively using this style is extremely important for your career no matter what field of Biology you intend to pursue. This is the style used to publish written works in the fields of Medicine, Veterinary Science, Dental Science, Cell Biology, Molecular Biology, Genetics, Botany, Ecology, Conservation Biology, and a host of other disciplines. This is the style that journal editors, and perhaps your employer, will insist upon. The best way to learn this style is to use it in writing and read it as it appears in the literature. In this course, you will practice the scientific writing style by using it to prepare your laboratory assignments. In fact, one of your first assignments is to obtain (from the library or from an electronic source, see below) a journal article of your choice, from a journal of your choice (any field or discipline of the biological sciences), to use as an example and to attempt to emulate when you prepare your lab assignments. You will also be given a specific example to follow (see the Appendices). Your ability to communicate complex scientific data and ideas (and your grade in this class) will improve dramatically if you carefully refer to these guidelines and the examples before your prepare each report. The Scientific Paper. A scientific paper is a written report describing new research results. In its prepublication form it is called a manuscript. That is, you write, and edit, and eventually send to the editor to be published, a manuscript. After it is published, it is generally referred to as an article. The scientific paper is a contribution to the scientific community at large, and is written as such. If not presenting new research results, a scientific paper may present a compilation or summary of previously published ideas. This type of article is called a review. Papers reporting new and original results are called "primary" sources, and reviews are considered "secondary" sources. An easy way to distinguish primary from

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secondary literature is to determine whether the authors actually performed the experiments they describe. If they did, the article is primary literature. Even though the scientific paper is written for scientists, it must be accessible to a wide audience of scientists from different fields. Even when directed to a very narrow group of biologists (e.g., cell biologists, or ecologists), different individuals in those fields have different knowledge and specialties, It is therefore important that the paper contain enough background information to prepare the reader for the material that follows. Not only must it introduce the reader to subject material, it also must integrate the work into the wider context of other papers in the field, past and present. It is often desirable, and useful for the reader, to suggest possibilities for future research, new lines of inquiry, and potential experiments to perform. The format used for scientific publication has been defined by a centuries-old heritage of tradition and practices based on scientific requirements as well as professional exchanges between scientists and printing and publishing services. According to this format, a scientific paper should have, in the order listed, the following sections: a Title Page, Abstract, Introduction, Materials and Methods, Results, Discussion, and References (or Literature Cited). Figure Legends (or Captions) are group together on their own page(s), and Tables and the Figures themselves should each appear on their own separate page. Your manuscripts (and Biol 350 Lab Assignments) should strictly adhere to this format. The ultimate source of instructions on how to prepare a manuscript for a given journal is printed once (or twice) yearly by the journal itself. This is the "Instructions for Authors" or "Guide for Contributors" or other similarly named set of instructions that is usually published in the January 1st issue of the journal. Though each manuscript contains a standard sequence of Title Page, Abstract, Introduction, etc..., this is not the order that most scientists write their papers. It is usually easier to write the Materials and Methods section first (theoretically, this can be done even before the data are analyzed), followed by the Results, Introduction, and Discussion. The Abstract is always written last so that it accurately reflects the finished work. This method will probably work for you, too. You can jot down notes concerning the basic sequence of events as you are doing the experiment. Homework Assignment. 1) Examine the "before" example provided on your instructor web site. These represents a "real life" example of a paper that was sent to a journal and subsequently published (in this case in the journal Comparative Biochemistry and Physiology). The "before" paper is called a manuscript and is what was mailed to the editor of the journal. You may be surprised at the simplicity of the manuscript, and how it resembles something you could easily prepare (i.e., no fancy typesetting of columns, figures not imbedded in text, etc...). Creating basic manuscripts based on your laboratory work that are precisely correct in format is a major goal of this course. 2) Find a suitable article of your choice from a respected, peer reviewed journal (also of your choice) and use it as a model for your own writing. "Suitable" means that it has all the sections mentioned above, i.e., Introduction, Methods and Materials,

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etc.... Bring this article to class with you the first week and keep it handy to use as a general guide throughout the semester. Writing the Manuscript: General Considerations. A manuscript that you intend to send to an editor must follow the journal's format precisely. You will give a very bad impression of yourself if you send in something that does not conform. The Editor will probably simply mail it back to you without comment (the ultimate publishing ignominy). Carefully read and follow the guidelines presented below. Though there are few rules concerning scientific writing that can be "written in stone", the following are good to keep in mind. · Write clearly and be brief. Scientific literature must be concise, terse, succinct. · Avoid vernacular (e.g., slang) and jargon as much as possible. When you must use it, explain it on its first use, if possible. · Always define abbreviations the first time you use them. Use the full length term first with the abbreviation in parentheses afterwards, never the other way around. E.g., "The concentration of docosahexaeneoic acid (DHA) was..." and not "The concentration of DHA (docosahexaeneoic acid) was...". · Avoid the use of phrases like "It can be seen that...", "It is clear that...", "We concluded..." and "We can see...". · Never refer to the Instructor of the class. Shun phrases like "The instructor prepared...", "Dr. Smith gave us..." and "We obtained from the instructor...". You will never do this in a real article. · Verb tense changes between sections; past tense is usually used for describing procedures (i.e., Materials and Methods), and present tense is used to describe results (Results) and conclusions (Discussion). However, be careful not to change tense within a section. · Include headings for each section of your paper. Follow the "Instructions for Authors", your guide article, and the examples in the Appendices. · In scientific writing quotations are extremely rare. Avoid using them. Instead, paraphrase the passage you wish to mention (in your own words) then cite the source. · Do not use the style of a cook book. Avoid phrases like "Add 3 mL of reagent A then mix...", "Place six tubes in a rack and label them....". You job is not to tell the reader what they are to do, but rather what you did. · Never start a sentence with a digit, always spell the number out if it is the first word of a sentence. For example use "Five mLs of reagent A....." rather than "5 mLs reagent A...". Finally, it well known by even the best authors that the first version of a manuscript is rarely flawless. Most of us struggle to different degrees to prepare our papers, and they usually require several revisions before we are satisfied with them. You will almost certainly find this to be true. Good scientific writing takes years of practice, and the more you do it, the more you will appreciate this fact. Do not be frustrated or feel

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defeated when your paper is returned to you with many red markings on it. We all have to go through this when we write. Do not take it personally; the comments are not attacks, they are editorial comments intended to help you become a better writer. The Components of a Scientific Paper Title Page. The Title of the paper is placed on the Title Page. It should accurately reflect the contents, emphasis, and perhaps the conclusions of the paper. Keep the title short and effective. Use the fewest possible words that straightforwardly describe the work. Avoid using the phrases "Results of...", "Investigations of ...", "A study of ...", "Observations on ...", etc. Journals place inviolable length restrictions on titles. Those limits are different for different journals so check the "Instructions for Authors" to be certain. Indexing and abstracting services, which are vital to your work reaching a wide audience who might find it interesting, depend heavily on the accuracy of the title. They use it to identify keywords that are then placed in large databases and used in computer searching. A poorly composed title may prevent your work from reaching those who would find it most interesting and useful. On the line(s) below the title should appear the names of all the authors. Below that appear the author's affiliations (i.e., their institutions, departments, etc...) and other contact information (address, phone number, e-mail address, etc...). To do this correctly, carefully follow the guidelines provided in the Instructions for Authors for the journal to which you are sending your manuscript. Requirements for Biol 350 Lab Reports. Give your report a Title Page consistent with the guidelines presented above. For this course, we will use the Instructions to Authors for the Journal of Cell Biology (www.jcb.org). These guidelines are available at the JCB web site, and are reproduced, in part, in Appendix 2 of this manual. (One exception to the Instructions: do not give your real phone number and home address on your lab reports! Use the Department of Biological Sciences as your address for correspondence and make up a phone number.) Abstract Though the abstract appears near the beginning of the manuscript, you should consider writing it last so that it accurately reflects the content of the manuscript. The abstract should state the problem, the principal objectives, and the scope of the investigation. Do not repeat the title. Include a brief statement of the methods used, but do not go into detail. Most importantly, the abstract should concisely summarize the results and principal conclusions of the paper. An effective way to write an abstract is to condense each section of the paper (i.e., Introduction, Materials and Methods, Results, and Discussion) into one or two sentences each. A well prepared abstract will allow readers to quickly identify the basic content of the paper so they can decide whether to read it in its entirety. Make it interesting! Journals generally limit the size of the abstract to between 100-300 words (depending on the journal), so it must be concise as well as accurate. Compose the

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abstract in as few words as will do the job well; if 100 words produces an excellent abstract, there is no need to use 200. The abstract and the title must not only accurately reflect what is in the manuscript, but they must be self-contained because they are published separately from the paper in abstracting services (e.g., Medline, Current Contents, Biological Abstracts). Exclude all references to tables and figures and do not include literature citations. Also do not include acronyms and abbreviations except for very common ones like DNA, RNA, ATP, etc..., even though they are defined elsewhere in the manuscript. Sentences such as "The effects of A on B are discussed in detail" and "The implications of these results are discussed" are useless. In general: · Try to include one to three sentences as each "section" of your paper (i.e., two sentences that serve as introduction, two for methods, two for results and two for conclusions). · Do not include literature citations in your abstract. · Do not include acronyms, spell everything out. Requirements for Biol 350 lab Reports. An abstract is not required for your Lab Reports. (But save these instructions for future use!) Introduction The introduction should be one to three paragraphs long. It should describe the background material that lead to your hypothesis, the hypothesis itself, and, and the reasons for doing the study. Also indicate how your work relates to published work and how it is different from those other works. Begin the introduction by describing the problem at hand and introducing the reader to the available literature. A well guided description of the problem and the pertinent literature is always preferred over a rambling and poorly focused stream of facts. Always use prose and avoid using a series of pointwise statements. Demonstrate an understanding of the relevant issues in the field and indicate how your study complements this information. After providing the background, clearly state the objective of the work. The introduction can finish with a statement of objectives or with a brief statement of the major findings of the work. It is important that the reader finish the introduction with a clear idea of where the manuscript is heading sos they can follow the development of the data you are about to present. Requirements for Biol 350 Lab Reports. The Introduction should comprise one to three paragraphs (rarely more). Try to accomplish the objectives listed above in one page or less. The introduction, along with the discussion, will contain the majority of your references. Materials and Methods This section can also be called "Experimental Methods". The main purpose of this section is to provide enough information about your procedures and methodologies so that a reasonably competent researcher can reproduce your results. Your results must be reproducible, and you have an obligation to provide a basis for repetition of your work by others. Write this section in the past tense as a description of the experiments you carried out.

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If desired, the Materials and Methods section can be divided into sub-sections by including sub-headings. For example, there may be subsections titled "Isolation of mitochondria", "Ion exchange chromatography", "Enzyme assay procedure", "Statistics", etc... Check papers published in the journal to determine if these are allowed. Include brief reference to equipment used and always completely describe all reagents and solutions used. Equipment obtained from commercial suppliers should be described exactly (e.g., Philips electron microscope, Model LB; Hewlett-Packard LS-55 Luminescence Spectrometer). Sources of materials should be given because there are often slight differences in quality or properties between suppliers (e.g., ...alcohol dehydrogenase was from Sigma Chemical Company, [St. Louis, MO]; bovine serum was from Hyclone, [Provo, UT]). The method used to prepare apparatus and reagents should be stated exactly, or a reference to a recipe used in a published paper should be given. No one will be able to repeat your experiments if you do not provide the recipe for the solutions you used. If relevant, you should also state any relevant conditions used when the test was performed (e.g., temperature, light, season, time of day, etc...). It is unnecessary to include descriptions of procedures that are familiar to most scientists. For example, its is not necessary to say "We added 1 g of succinate to 95 ml of distilled water, then brought the mixture up to 100 ml". Instead say "We prepared a 0.2 M solution of succinate" or just "0.2 M succinate was used.....". It is also unnecessary to include such trivia as the number of the tube or gel lane a sample was placed in, or the number of total tubes or gel lanes unless the procedure requires a specific knowledge of these facts. For example, never say "sample number 1 was loaded into lane number 1," or "six tubes were labeled 1 through 6". You may choose to present your methods chronologically, but this not a requirement. Often, it is better to describe related methods in the same section regardless of the order that they were used. Be precise in describing measurements and be sure to indicate which error of measurement (e.g., standard deviation, standard error, range) you are using where appropriate. Ordinary statistical methods can be used without comment, but advanced or unusual methods should be accompanied by a literature citation. Under some circumstances, or for some journals, it will be necessary to justify the statistical techniques you used. Requirements for Biol 350 Lab Reports. Follow the directions above. This section should be between one and two pages in length. Look over the Materials and Methods sections of the paper you obtained from Homework Assignment 2 above and the "before" and "after" manuscript in the Appendix of this manual to get a sense of how to properly compose this section. Results This section is where you present the outcomes of your experiments. Present your data in abridged form (i.e., pre-processed and condensed) and guide the reader through clear and simple descriptions of important trends and differences. Always remain objective in your presentation. Your results represent the new knowledge that you are contributing to science, and it is important that you remain detached and unemotional. Avoid statements expressing satisfaction or disappointment. Use rational arguments and logical flow to convince your reader that your arguments are sound. If they are confused

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by your presentation, or do not follow your arguments, they may not accept your conclusions. Before you start writing be sure you have thoroughly studied your data and that you are clear about what information you want to convey to your reader. Once you have decided on the points you wish to make, design your presentation to highlight those points as clearly as possible. Some simple results can be presented in the text. More complex data will need to be presented in either a Table or a Figure. It will be up to you to decide which format (Table or Figure) is most appropriate for a particular data set. Use the format you believe most effectively communicates the ideas you wish to convey. If the exact number for each data point is important, then a table is probably warranted. When a trend or pattern between data points is important then use a table. Tables and figures should be titled and captioned in such a way that they are understandable on their own, without reference to the text (but the placement of titles and captions are different for table and figures, see the guidelines below). Both figures and tables should include only the data that pertains to the points you are making when referring to that table. Don't include "unused" data or trivial or distracting information. If it is important data, present it, otherwise do not include it (no matter how hard you worked to get it!). The results section should be short and sweet. Do not babble. Be terse. For example, instead of "It is clearly evident from Fig. 1 that cell growth increased with the amount of serum added to the medium." say "Cell growth increased with added serum (Fig. 1)". Avoid giving long lists of results without providing a brief evaluation, explanation, or interpretation (though most of this will be done in the Discussion). For example, the series of phrases "Serum level significantly affected growth (Table 1). pH significantly affected growth (Table 2). Light levels had no significant effect on cell growth (Table 3)" leave the reader thinking "so what?". Develop each idea and describe the effect. How did the levels of the independent variable differ? However, your results section should only include direct biological interpretations ("the cells grew", "the enzyme was active"). Save indirect interpretations ("the increased activity of this enzyme may help these cells survive when...") for the Discussion section. Also, by convention the first table that is referred to in the results section is "Table 1" and the first figure referred to, regardless of whether it falls before or after Table 1, is called "Figure 1". It is a good idea number your tables and figures only after you have written your Results section. This helps ensure that the flow of your data and results are logical and not dictated by the order of the experiments. Requirements for Biol 350 Lab Reports. We will be particularly interested in proper presentation practices. Include enough text in this section to guide your reader through the tables and figures you present. The text part of the Results section can be a short as one paragraph. Figures and Tables make up the bulk of this section (but these are presented at the end of the text, see below) Preparation of Tables and Figures General considerations · Present data in either a figure, a table, or in the text, but never present the same data twice. · In the text of the paper always refer to every table and figure included.

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· Do not present data in the form of tables or figures if it can easily be stated in a sentence or two of text. · Do not title your tables and figures, but be sure to give them legends. Most journals require you to present figure legends together on a separate page. Figure legends should not be printed on the same page on which the figure appears. (This may seem strange, but the legends and the figures are treated differently by the typesetter, so it is easier to have them on separate pages.) Each figure should appear on its own page, and the primary author's name and figure number should be printed (or handwritten in) on the very bottom right hand corner of the page. See the example manuscript in the Appendix. Tables · Tables, which are treated as text by the typesetter, should have at their head (i.e., at the top) the table number and a caption. The caption should be brief, but should allow the figure to stand alone. The reader should be able to understand it without referring to the text. · Include appropriate and precise column headings. · Organize tables so that like elements read down, not across. · Do not include columns of data that contain the same value throughout. If the value is important to the table include it in the caption or as a footnote to the table. · Tables should contain horizontal rules (lines) only. Never use vertical rules in a table. Figures · Always include, on a separate page, the captions (some journals call them legends) describing each figure. Captions (legends) should be very brief, yet still contain enough information for the reader to understand the figure without reference to the text. See the sample manuscript in the Appendix. · Label both axes with a brief but informative title. Always include units of measurement on the axes! · Do not fill the entire page with a graph leaving little room for axis numeration and axis titles. · Match the extent of the axes to the data in the figure. For example, if the data range between 0 and 27, the axis should extend to 30, 35 or maybe 40 or 50, but not to 100 or 1000. · Each figure should be placed on a separate page (but they do not need to cover the entire page!). · Avoid grid lines on figures unless they are specifically required to make a point. · Make all figures in black and white. Avoid the use of color (which adds enormously to the cost of publication). The background should be white. · Proportion the aspect ratio of your figures appropriately. Avoid "squashed" (yaxis too short) or tall (x-axis too short) figures. · Do not place figure legends on the graph itself. Identify symbols in the text of the figure legend.

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· Be careful not to allow graphics software to dictate to you how your figure should appear. For example, Excel automatically adds grid lines to all graphs it produces and makes the background gray. Be sure to remove the grid lines, unless you feel they are necessary, and to change the background to white. Other Graphics (SDS-PAGE gels, western blots, etc...) · Label the lanes of gels. Use the sample name or number or label the lanes with numbers that are then explained in the Figure Legend. · Use arrows to indicate bands that are discussed in the text. Requirements for Biol 350 Lab Reports. Meticulously follow the directions given above. Discussion The Discussion is where you explain and interpret your results and interconnect them with the background material presented in the Introduction. Conclusions are presented and discussed in this section. Base your conclusions on the evidence you presented in the Results section. Address the objectives of the study in this section and discuss the significance of the results. Explain your new findings in the light of previous work in the literature. What new principles have been established or previous ones reinforced? What generalizations can be made of this or of other systems? How do your findings compare to those in the literature? Was your hypothesis supported or challenged? Were the major hypotheses in the field supported or challenged? Are there any theoretical or practical implications of your work? Be careful that while addressing these questions you base your statements on the data you presented in the Results section. Do not extend your conclusions beyond those that are directly supported by your data. You may choose to speculate here, but do not let it form the bulk of the discussion. You may choose to include suggestions for future research, or disclaimers and explanations of methodological errors made during the course of the experiment. However, NEVER say things lke: "Our experiment failed because...' or "Our experiment did not work because.,..". If your experiment failed, why are you writing about it? No..., your experiment worked, just not in the way you intended or expected. Finally, don't leave the reader thinking "So what?". End the discussion with a short summary or conclusion regarding the significance of the work. Requirements for Biol 350 Lab Reports. For our purposes, aim for a Discussion one page or two in length. It is important to include references in this section. References When you use data, ideas or other information from the literature, you must cite the source. The resulting list of references is then included at the end of the paper in the "Literature Cited" or "References" section. The format for references, and how to cite the references in the body of the paper vary between journals. For this class, you will use the style found in the Journal of Cell Biology (the Instructions to Authors for JCB can be found at http://www.jcb.org/misc/ifora.shtml). (Use this format regardless of the

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"Instructions for Authors" you copied in the library or printed from a web site. Everyone in this course will use this format for their References section.) In the body of the text, references should be cited using the author's name and date, e.g. (Smith, 2003) or (Smith and Jones, 2004). In cases of multiple citations, list them chronologically separated by semicolons: (Smith, 2001; Jones, 2002; Johnson and Brown, 2003; White et al., 2004). If there are more than three authors, use the first author's name followed by "et al.": (Smith et al., 2003). In the References section, references should be listed alphabetically by first author's last name. List all author's names (i.e., do not use "et al." in the References section). References should contain complete titles and inclusive page numbers. Abbreviate the names of journals as in the Serial Sources for the Biosis Data Base (available in the library), or as listed by PubMed (http://www.ncbi.nlm.nih.gov/entrez/query.fcgi/ OR ftp://ftp.ncbi.nih.gov/pubmed/J_Medline.txt ). Please note that URLs are unacceptable as references. The original paper to which the URL points or refers is acceptable if it has been published in a legitimate print journal. Requirements for Biol 350 Lab Reports. Follow the directions above. Lab reports for Biol 350 must have at least three references from the primary literature. Format for references: Journal Articles. Two authors: Yalow, R.S., and S.A. Berson. 1960. Immunoassay of endogenous plasma insulin in man. J. Clin. Invest. 39:1157-1175. More than two authors: Benditt, E.P., N. Ericksen, and R.H. Hanson. 1979. Amyloid protein SAA is an apoprotein of mouse plasma high density lipoprotein. Proc. Natl. Acad. Sci. USA. 76:4092-4096. Complete Books Myant, N.B. 1981. The Biology of Cholesterol and Related Steroids. Heinemann Medical Books, London. 882 pp. Articles in Books and Book Chapters Innerarity, T.L., D.Y. Hui, and R.W. Mahley. 1982. Hepatic apoprotein E (remnant) receptor. In Lipoproteins and Coronary Atherosclerosis. G. Noseda, C. Fragiacomo, R. Fumagalli, and R. Paoletti, editors. Elsevier/North Holland, Amsterdam. 173-181.

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These guidelines were prepared with the aid of a number of sources. You might find the following particularly useful. Davis, M. and G. Fry. 1996. Scientific Papers and Presentations. Academic Press, New York. 296 pp. R. Day. 1998. How to Write and Publish a Scientific Paper. Fifth Edition. Oryx Press, Philadelphia, PA. 275 pp. J. S. Dodd, ed. 1997. The ACS Style Guide: A Manual for Authors and Editors. Second Edition. American Chemical Society Press, Washington, DC. 460 pp. CBE Style Manual Committee. 1983. CBE Style Manual: A Guide for Authors, Editors, and Publishers in the Biological Sciences (Council of Biology Editors Style Manual). Fifth edition. Council of Biology Editors, Inc. Bethesda, Maryland. 324 pp. Hall, G. M. 2003. How to write a Paper. Third edition. BMJ Publishing Group. Anapolis Junction, Maryland. 176 pp. Huth, E. 1987. Medical Style and Format: An International Manual for Authors, Editors, and Publishers. Lippincott Williams & Wilkins, Philadelphia, Pensylvania. 355 pp.

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Chapter 2.

Pipetting and Creation of a Standard Curve.

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Objective Practice using micropipettors to create a standard curve. Use the standard curve to determine the protein concentration of an unknown. Introduction In the laboratory section of Cell Biology, you will frequently use micropipettors to transfer small volumes of liquids. You will also regularly create and use standard curves to analyze and assess data you have collected. Both of these techniques are very commonly encountered in the real world of science. This exercise is designed to familiarize you with them; you will use micropipettors to construct a standard curve. Micropipettors Pipettes are slender, graduated tubes, usually of glass or clear plastic, used to measure the volume of fluids or to transfer them between vessels. Micropipettors are mechanical devices used to accurately measure small volumes of liquids, usually less than 1 milliliter (mL). There are two basic types of micropipettors, fixed volume and adjustable volume. As the name implies, fixed volume micropipettors are preset at the factory to deliver a single volume every time they are used. Adjustable volume micropipettors, like the ones we use in the Cell Biology Laboratory, can be set to deliver a fairly wide range of volumes. These are manufactured to deliver different ranges of volume, e.g. one might deliver volumes in the range between 1 and 100 L and another volumes in the range between 100 and 1000 L. The ability to accurately measure and transfer small volumes of liquid is essential to your success in the Cell Biology Laboratory. In use, the micropipettors itself should never touch the liquid that is being transferred. Instead, a disposable tip is placed on the end of the device and the tip is submerged in the liquid. Never use a micropipettor without a tip. To avoid cross contamination between liquids, change the tip between each use. When in doubt, change the tip. Tips are color coded. In general, blue pipette tips are used with micropipettors with capacities of 200-1000 µl, yellow tips are used with micropipettors with capacities of 5-200 µl, and clear ("natural") tips are used with the smallest micropipettors, those with capacities of 0.5 to 10 L. Table 1 defines some units of volume that your will encounter in this course and in the scientific literature. How to operate a micropipettor. General considerations. · Handle these devices with care. Rough handling can cause them to lose there calibration, and therefore their accuracy. Dropping them can cause irreparable damage.

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Table 1. Units of volume. Unit abbreviation definition liter L milliliter mL 1/1000th of a liter (also equal to a cubic centimeter or "cc") microliter L 1/1000th of a mL nanoliter nL 1/1000th of a µL picoliter pL 1/1000th of a nL ________________________________________________________________________ · Never force an adjustment knob. If the knob becomes hard to turn, it is probably at the limit of its travel. Forcing it can cause irreparable damage. · Never use a micropipettor outside of its specified range. · Do not use micropipettors to measure strong acids, bases, or flammables. · For the greatest accuracy in volume measurement, select the smallest size pipette that will handle the volume you wish to transfer. Accuracy is reduced when an unnecessarily large pipette is used to measure small volumes. General operation. 1. Select the most appropriate sized micropipettor for the job at hand. 2. Put on the appropriate sized tip by tapping the micropipettor into the tip while the tip is still in its rack. Do not touch the tip. 3. Set the desired volume by turning the adjustment knob as shown in Figure 1. (Micropipettors from different manufacturers have different adjustment mechanisms. Check with your instructor if the micropipettor you are using does not look like the one in Figure 1.) _ _

Figure 1. Adjusting the desired volume on a micropipettor. 4. Figure 2 shows the proper operation of a micropipettor. Before putting the tip into the sample solution, depress the thumb knob to the first stop. 5. Immerse the tip approximately 3 mm into the sample solution (step 1).

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6. Slowly release the thumb knob to the initial position (step 2). Do not release it too quickly, the sample might cavitate and form a bubble, destroying the accuracy of your volume measurement. 7. Withdraw the tip from the sample solution. 8. Place the tip against the side wall of the receiving container (step 3). 9. Smoothly depress the thumb knob to the first stop (step 4), pause, then depress the knob to the second stop (step 5). 10. Remove the tip from the receiving container, and return knob to the initial position. Do not let the knob snap back. 12. Remove the disposable tip by firmly depressing the tip ejector knob (step 6). 13. Add as new tip and continue.

Figure 2. Operating the micropipettor.

Construction of a standard curve. An assay is a test or an evaluation. The term often refers to an experimental procedure used to estimate the amount or concentration of a substance in a sample. Such assays are often based on the a measurement of a physical or chemical property of one compound that changes in the presence of the compound of interest. The measured property might be fluorescence, the ability to form a precipitate, optical density, radioactivity, a color change, or something else. A standard curve is a graph of assay measurement (Y) as a function of known amounts or concentrations of the substance of interest (X). Known amounts or concentrations of the substance of interest are used to calibrate the assay. Standard curves can be linear or curved. Once a standard curve has been created, it can be used to determine the amount or concentration of the substance of interest in other samples (unknowns). The same assay is performed on the unknowns and the value of the measured property (Y) is recorded. The measured value of Y is then read across the graph from the ordinate (y-axis) to the standard curve, then read down to the intersection of the abscissa (x-axis). The amount or

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concentration of the substance of interest in the unknown sample is read off the abscissa. In this exercise you will measure the change in color of a reagent that is normally brown and turns blue in the presence of protein. You will test your skill in using the micropipettes by constructing a standard curve for determining the protein concentration of an unknown sample. If you use the pipettes correctly, the curve that you generate in this exercises will be a straight line with an r2 value of 1.00. r2 is a estimate of "goodness of fit" of the line to the data. Values of r2 range from 1 (perfect fit) to 0 (no fit, similar to a line drawn through data that have a pattern like a shotgun blast). The concentration values that you determine for your unknown should be within 10% of the actual value (which is known by your instructor). This exercise will not be graded and you will not be required to hand in a written report. However, the experience you gain in this exercise will be essential in future laboratory exercises, and there are (ungraded) homework problems to complete.

16

Procedure 1. Set the spectrophotometer to 595 nm. 2. Set up a rack of six test tubes and label them 1-6 with a marker. 3. To each tube add the indicated volumes of the solutions listed in the Table 1 below, then cover each tube with parafilm and mix by inverting slowly several times. The concentration of the protein stock solution ("protein sol" in the table) is 2mg/mL. Table 1. Reagent volumes for preparation of the protein standard curve. Tube water protein std assay concentration Absorbance595 number reagent (L) (L) (2 (mLs) mg/mL) 1 100 0 4.0 _____________ _____________ 2 95 5 4.0 _____________ _____________ 3 90 10 4.0 _____________ _____________ 4 80 20 4.0 _____________ _____________ _____________ 5 65 35 4.0 _____________ 6 0 100 of one 4.0 _____________ _____________ of the unknowns 4. Calculate the concentration of protein in each of tubes 1-5. What are the appropriate units for these concentrations? Units: _________________. 5. Insert each tube into the spectrophotometer (after wiping with a kimwipe to remove fingerprints) and measure the absorbance of the mixture at 595 nm. Record your data. 6. Using the Excel software on the computer at your station, plot absorbance versus protein concentration. Which values should be plotted on each axis? Y-axis: _____________ X-axis: _______________ 7. Use Excel to plot a linear regression through your data. Should this line be straight or curved? Does your regression line go through zero? _____ Should it? _____ 8. Use the software to determine the formula for your regression line. What is the formula for any line? What is the formula for your regression line? any line: _______________ my line:_______________________ 9. Use the software and the formula for your regression line to determine the concentration of protein in the "Unknown" sample. Concentration of Unknown ______.

17

Chapter 2. Homework Assignment. Produce standard curves from the following data and answer the questions that follow. 1. g protein/mL 0 5 10 15 20 25 30 35 Absorbance 0.000 0.087 0.176 0.272 0.368 0.428 0.465 0.480

a. What happens to the shape of the curve above 20 g protein? b. Can you suggest several reasons for this change in shape (i.e., what could have caused this in a "real" experiment)? c. Would you use this standard curve for your experiment? Why or why not? 2) mmol NADH 0 10 20 30 40 50 60 Absorbance 0.360 0.351 0.343 0.334 0.325 0.346 0.307

a. How is this assay fundamentally different from the one in question #1 above? (I.e., What is the basic, visual difference between these two assays?) b. What happened to the point at 50 mmol NADH? c. Would you use this standard curve for your experiment? Why or why not?

18

Chapter 3.

Cell Fractionation: Isolation of Mitochondria from Cauliflower.

___________________________________

Objectives: Fractionate cauliflower cells using differential centrifugation. Introduction An important concept in experimental cell biology is that of "fractionation". The term fractionate means to "separate or divide into component parts". In the context of cell biology, the term is applied to a very large number of techniques used to separate and isolate cell components. Eukaryotic cells contain many different organelles, each with distinct functional properties. These properties were discovered and elucidated, in many cases, by isolating the organelles from the cells in which they originated. Organelles can be removed from cells and separated from one another using a process called "cell fractionation". Furthermore, each organelle can be further fractionated into its constituent parts, for instance, its unique set of proteins. The results of such fractionation procedures are a series of fractions, each one, ideally, containing only one, highly purified organelle or protein. For instance, a liver cell could be used to produce one fraction containing only nuclei, another containing only mitochondria, another containing only lysosomes, etc.... A fraction of purified mitochondria could then be used to fractionate mitochondrial proteins, for example the enzymes of the tricarboxylic acid cycle, which could then be separated and studied individually. The fractionation of cells into organelles always begins with the disruption of the cells to release their contents into a benign buffer. The methods used for forming this initial "cell homogenate" depend on the organelles that are sought and on the nature of the cell containing them. For example, plant cells, which are surrounded by a cell wall, are frequently disrupted by grinding the tissue with a mortar and pestle in the buffer and in the presence of an abrasive. Animal tissues and cells may be similarly treated in less substantial, all glass mortar and pestle systems without using abrasives. They can also be effectively disrupted by repeated freeze-thaw cycling, cavitation (explosive decompression of cells saturated with a compressed gas), sonication (high frequency sound waves), forcing the cells through filters with small pore size, osmotic lysis, and a number of other means. Once a suitable cell homogenate has been prepared, the organelles may be separated from one another using an equally wide variety of methods. Most any property of the organelles can be used as a basis for their separation, including differences in size, density, charge, and other properties. Fractionation methods include size exclusion chromatography, affinity chromatography, immunoprecipitation, two-polymer partition, and other, fairly exotic, techniques. By far the two most popular methods, probably because they are the most flexible and effective, use gravity to separate organelles based

19

on their size and density. These tow methods are differential centrifugation and density gradient centrifugation. In differential centrifugation the organelles are allowed to settle to the bottom of the tube while the tube is exposed to increasing, incremental steps in gravity (Figure 1). The increase in gravity is produced by spinning the tubes in a centrifuge, much the same way that astronauts and pilots are trained to tolerate high g (gravity) forces. Increasing the rotation rate of the rotor causes an increase in the gravitation force experienced by the homogenate in the tube. The gravitational force created by a spinning centrifuge rotor can be great. Particles that would ordinarily remain in suspension indefinitely at 1 g will fall out of suspension and form a pellet at the bottom of the tube at higher g forces. The relative centrifugal force produced by a spinning rotor can be calculated by the equation: RCF = 1.119 x 10 -5 (rpm2) r where rpm is the revolutions per minute of the rotor and r is the distance (in cm) of the particle from the axis of rotation. In practice, the radius used is the distance from the center of the axis of rotation to the middle of the centrifuge tube. The gravitational forces

Figure 1. Differential centrifugation.

created even at low centrifugation speeds are very large compared to that on earth (e.g., 600 g) but are nevertheless only able to pellet very large or dense particles At high speeds, the gravitational forces created can be enormous (e.g. as much as 300,000 g). At these speeds, most particles will fall out of suspension and only very small, highly soluble molecules will remain in solution. Thus, once a cell homogenate has been prepared, the next step in the cell fractionation process is to subject it to a relatively low speed spin (50-200 g) for a few minutes (2-15 min). Large, dense and/or heavy particles, such as unbroken cells, bits of

20

undisrupted tissue, connective tissue, and abrasives (if present) tend to sediment quickly during this initial spin. After centrifugation, the pellet containing the sedimented particles is separated from the supernatant over it by aspiration or by carefully decanting. The supernatant is then returned to the centrifuge and spun again at a higher speed and for a longer duration. During this second spin, the next heaviest particle form the pellet, and the resulting supernatant is recovered and spun again. This process can be repeated as many times as desired. The final steps involve very high speeds (100,000 ­ 300,000 g or more) coupled with very long exposure times (3-24 hours). In practice, four or five separate centrifugation steps are employed. This constant sedimentation of particles base on time and exposure to increasing gravity is termed "velocity sedimentation". Density gradient centrifugation is similar, but this method employs a density gradient made of glucose, sodium chloride, cesium chloride, or a solute specifically designed for density separations such as Percoll® or Ficoll®. As the tube is spun the gradient material establishes a density gradient. The organelles then settle at their buoyant density and stop migrating in the tube. This is called "equilibrium sedimentation". The tube is emptied by upward displacement using a saturated sucrose solution or the bands are aspirated off individually. These two methods have proven to be very effective methods of cell fractionation.

Table 1. Enzymes commonly used to identify organelles. ________________________________________________________________________ Organelle Enzyme(s) ________________________________________________________________________ Nuclei DNA, Histones, DNA polymerase Mitochondria Succinic dehydrogenase, Cytochrome oxidase Lysosomes Acid phosphatase, Other acid hydrolases Peroxisomes Catalase, Urease Plasma Membranes 5'-Nucleotidase, Na+/K+-ATPase ER Glucose-6-phosphatase, Nucleoside diphosphatase Golgi Glycosyl transferases Cytosol Amino acyl tRNA synthetases, Glycolytic enzymes ________________________________________________________________________

Once the fractions have been recovered, it is necessary to establish distribution of each organelle (or the organelle of interest) among those fractions. Rarely will a fractionation result in the perfect separation of organelles. More likely, a given fraction will be enriched with a certain organelle that occurs, to some extent, in several fractions. Establishing this distribution can be accomplished by a number of means. For example, one could observe a sample of each fraction under a microscope and assess which organelles were present. However, this would require an electron microscope for most organelles, and processing the samples for electron microscopy would be tedious and time consuming. A somewhat easier method that enjoys widespread use and acceptance is one that measures enzyme activities. The cell fractions are measured for the presence

21

of an enzyme unique to each organelle. For example, succinate dehydrogenase and cytochrome c oxidase are found nowhere in the cell but the mitochondrial inner membrane. Similarly, Na+/K+-ATPase and 5'-nucleotidase occur only in the plasma membrane. Examples of other such "marker enzymes" are listed in Table 1. Thus one could, for example, measure the activity of succinate dehydrogenase in each fraction. The fraction containing the highest activity is the one with the most mitochondria. A similar assessment could be carried out for any number of enzymes, the enzymatic make-up of each fraction could be well characterized, and in this way the distribution of the originating organelles can be established. The centrifugation steps can then be modified to increase the proportion of the organelle of interest in a particular fraction. In this lab you are going to use differential centrifugation to fractionate cauliflower cells. You will determine the distribution of mitochondria in the resulting fractions by assaying for the activity of the mitochondrial enzyme succinate dehydrogenase.

22

Procedure. Note: Be sure to keep all cell preparations on ice, at all times. 1. Cut off several florets from a head of cauliflower. Clean a razor blade with ethanol and use it to shave off the outer 3-5 mm of the floret into a large weigh boat. 2. Weigh the tissue (aim for approximately 25 g). 3. Place the tissue in a cold (get it from the freezer) ceramic mortar, add 20 mL of icecold isolation buffer, and 6 g of cold abrasive (sea sand). 4. Grind the tissue until it is a fairly smooth paste (approximately the consistency of ground garlic found in jars at the grocery store). 5. Add an additional 20 mL of isolation buffer and continue to grind for another minute or two. 6. Place four layers of cheesecloth in a funnel and strain the homogenate into a 50 mL centrifuge tube supported in a beaker of ice. 7. Wash the mortar with an additional 5 mL of isolation buffer and pass this through the cheesecloth too. Squeeze the cheesecloth to collect any remaining homogenate. 8. Centrifuge the homogenate at 600 g (Baxter Megafuge, 1800 rpm) for 10 minutes at 4°C. Label your tube then balance it with that of another group. Place the two balanced tubes in opposite positions in the rotor. 9. Carefully decant the postnuclear supernatant into a clean "Oak Ridge" tube. Discard the pellet. (Note: Oak Ridge tubes are not disposable. Do not throw these away, save them to be washed!) 10. Centrifuge the postnuclear supernatant at 12,000 g for 30 minutes at 4°C (in the SS34 rotor). (While the supernatant is spinning, answer these questions: Why did you just throw away the pellet! What was in it?) 11. Decant the postmitochondrial supernatant into a graduated cylinder. Measure and record its volume. Volume of postmitochondrial supernatant _______________________. 12. Mix the supernatant well then transfer approximately 5 mL of it to a clean 15 mL centrifuge tube. Label the tube "postmito supernatant" and put it in a rack in the freezer. Discard the remaining supernatant.

23

13. Add 5 mL of cold assay buffer (NOT isolation buffer) to the mitochondrial pellet remaining in the tube. Resuspend the mitochondrial pellet by vortexing vigorously. (Why is it very important that the pellet be resuspended to homogeneity?) Measure and record the volume of the mitochondrial pellet. Volume of mitochondrial pellet ______________________. 14. Transfer the suspension of (putative) mitochondria to a new 15 mL centrifuge tube and label the tube "mito". Place the tube in the freezer next to your other tube.

Media Compositions Isolation buffer (0.3 M D-mannitol, 0.02 M phosphate buffer, pH 7.2). Assay buffer (0.3 M D- mannitol, 0.01 M KCl, 0.005 M MgCl2, 0.02 M phosphate buffer, pH 7.2). This laboratory exercise was adapted and modified from Exercises in Cell Biology for the Undergraduate Laboratory, a project of the American Society for Cell Biology Education Committee, 1992. (https://www.ascb.org/publications/exercises.html) References Ahern, H. 1997. Introduction to Experimental Cell Biology. McGraw-Hill Companies, Inc., Dubuque, Iowa. Crupper, S.S., and J. T. Dawson. 1999. A novel approach to the isolation of plant mitochondria in the introductory college biology laboratory. Bioscene 25: 7-10. Lambowitz, AM. 1979. Preparation and analysis of mitochondrial ribosomes. Methods Enzymol. 59:421-33. Zhao, J., T. Onduka, J .Y. Kinoshita, M. Honsho, T. Kinoshita, K. Shimazaki, and A. Ito. 2003. Dual subcellular distribution of cytochrome b5 in plant, cauliflower, cells. J Biochem (Tokyo) 133:115-21.

24

Chapter 4.

Cell Fractionation: Assay of Mitochondrial Enzyme Activity.

___________________________________

Objective Determine the distribution of mitochondria in the subcellular fractions prepared previously by assaying for the activity of the inner mitochondrial membrane enzyme succinate dehydrogenase. Related reading in text: p. 326-330. Introduction In this experiment, you will determine the distribution of mitochondria (Fig. 1) in the subcellular fractions you collected last week by assaying for the activity of succinate dehydrogenase (SDH). SDH is one of the nine enzymes that catalyze the steps of the tricarboxylic acid cycle (citric acid cycle, Kreb's cycle, Fig. 2 and see Figure 14-8 on p. 408 of your text).

Figure 1. A mitochondrion.

and see Figure 14-8 on p. 408 of your text). However, unlike the other enzymes of the TCA cycle, SDH is not a soluble enzyme found in the mitochondrial matrix, but is instead bound to the inner mitochondrial membrane (Fig. 2). (Why is this fact critically important for your experiment? Hint: Why did we choose to assay this particular enzyme and not one of the other eight involved in the TCA cycle?) SDH catalyses the oxidation of succinate to fumarate (succinate + FAD Fumarate + FADH2, Fig. 3). FAD accepts the electrons liberated in that reaction and is reduced to FADH2. FADH2 then passes the electrons to ubiquinone, the first of several electron acceptors that constitute the electron transport chain. You will measure the activity of succinate dehydrogenase by replacing ubiquinone with an artificial electron acceptor, 2,6-dichlorophenolindophenol (DCIP). The oxidized form of DCIP accepts electrons from FADH2, and this reduction causes DCIP to change color from deep blue to colorless. This change can be measured spectrophotometrically. To ensure that the

25

electrons are passed to the dye and are not funneled into the electron transport chain, the poison sodium azide is added to the reaction to block the final transfer of electrons from cytochrome a3 to oxygen. This effectively bottlenecks the electron transport chain, preventing FADH2 from passing along its electrons to ubiquinone. The basic procedure is to mix samples of fractions to be tested (enzyme) with an assay mixture containing substrate (succinate), cyanide and DCIP. The rate of the disappearance of the blue color is proportional to the concentration of enzyme. The change in absorbance of the mixture is measured as a function of time and the molar amount of DCIP reduced per minute is calculated.

Figure 2. The TCA cycle. There are two general types of enzyme assays: kinetic assays and fixed time assays. Kinetic assays follow either the consumption of a reactant or the formation of a product over time. These methods are commonly used when one of the products or reactants absorbs at a specific wavelength, fluoresces, or possesses some other property that can be readily monitored. In fixed time assays the amount of reactant consumed or product formed is assessed after the reaction has been allowed to proceed for a fixed interval. This method is common in instances where the species being monitored can only be detected after some chemical, physical or other type of processing. In fixed time assays it is important to establish that the reaction occurs linearly over the fixed time. A

26

kinetic assay must be performed to establish this fact. For both types of assays it is important that the reaction rate be proportional to the amount of enzyme added.

SDH HOOC-CH2-CH2-COOH + FAD Succinate HOOC-CH=CH-COOH + FADH2 Fumarate

Figure 3. The oxidation of succinate to fumarate is catalyzed by SDH.

27

Procedure. A. Preparation of a DCIP standard curve. 1. Turn on the spectrophotometer and set the wavelength to 600 nm. 2. Obtain 6 13 x 100 mm tubes and a tube rack. Label the tubes 1 through 6. 3. Prepare dilutions of 50 M DCIP solution as shown in the Table 1 below: Table 1. Reagent quantities for preparation of the DCIP standard curve. Tube # 1 2 3 4 5 6 Assay Buffer (ml) 5.0 4.8 4.6 4.4 4.2 4.0 DCIP (ml) --0.2 0.4 0.6 0.8 1.0 Concentration (M) ________________ ________________ ________________ ________________ ________________ ________________ Absorbance600 (AU) ________________ ________________ ________________ ________________ ________________ ________________

4. Cover each tube with a small piece of parafilm and invert several times to mix. 5. Read and record the absorbance at 600 nm (A600) of each tube. Use the tube containing 0 µM DCIP (tube #1) as a blank. 6. Use Excel to construct a DCIP standard curve. Plot the A600 of each tube against its DCIP concentration.

B) Succinate Dehydrogenase Assay. Note: Sodium Azide is a poison and should be handled carefully! 1. Obtain 6 13 x 100 mm tubes and a tube rack. Label the tubes 1 through 6. 2. To each tube, add assay buffer, sodium azide, 0.2 M succinate, and 50 m DCIP, as shown in Table 2 below. Cover each tube with parafilm and invert several times to mix thoroughly. Note: Before mixing the 6 solutions indicated below, read steps 3-6 below. They contain important information on the timing of this procedure that must be followed to obtain accurate results. Also, vortex the mitochondrial

28

suspension and postmitochondrial supernatant before using to ensure that they are homogenous (Why is this important?) and be sure to keep them cold throughout the procedures. Table 2. Reagent mixtures for use in the assay of SDH. Tube Assay Buffer (mL) 3.5 3.0 3.5 3.0 4.0 3.5 Sodium Azide (mL) 0.5 0.5 0.5 0.5 0.5 0.5 Succinate (mL) 0.5 0.5 0.5 0.5 0.5 0.5 DCIP (mL) Mito. (mL) 0.5 0.5 --------Postmito. Super. (mL) ----0.5 0.5 -----

1 2 3 4 5 6

--0.5 --0.5 --0.5

3. Add 0.5 ml of mitochondrial suspension to tubes 1 and 2, and note the time (this is time = 0). Immediately following the addition of the mitochondria, mix the tubes thoroughly and read the A600 of tube 1 (blank) and tube 2. Then place the tubes in a test tube rack at room temperature. 4. Add 0.5ml of post-mitochondrial supernatant to tubes 3 and 4, again noting the time. Immediately mix the tubes and read the A600 of tube 3 (blank) and tube 4. Then place the tubes in a test tube rack at room temperature. 5. Mix tubes 5 and 6 thoroughly, and read the A600 of tube 5 (blank) and tube 6. Then place the tubes in a test tube rack at room temperature. 6. At 5 minute intervals read the A600 of tubes 2 and 4. Remember to use the correct blank to adjust the spectrophotometer before each reading. Continue to collect absorbance readings for 30 minutes. Record your data in Table 3. (Tube #6 is the control tube and only needs to be read at time = 0 and time = 30 min. The absorbance should not change since DCIP is not being reduced. But...does it?).

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Table 3. SDH activity data. Time (min) 0 5 10 15 20 25 30 Abs600 Tube #2 _______ _______ _______ _______ _______ _______ _______ Conc. Tube #2 ________ ________ ________ ________ ________ ________ ________ Abs600 Tube #4 ________ ________ ________ ________ ________ ________ ________ Conc. Tube #4 ________ ________ ________ ________ ________ ________ ________ Abs600 Tube #6 ________ ___···___ ___···___ ___···___ ___···___ ___···___ ________ Conc. Tube #6 ________ ___···___ ___···___ ___···___ ___···___ ___···___ ________

7. Use the regression equation established in Part A above to convert the absorbance data just collected (Table 3) into concentrations.. 8. Construct a plot concentration of reduced DCIP in each tube against time for the mitochondrial solution and for the postmitochondrial supernatant (plot both lines on the same graph). Compare the two lines. 9. Calculate and compare the micromoles of DCIP reduced per minute for each of the tubes (I.e., the slopes of the lines). What are the units of the slopes of these two lines? 10. Save your data to use next week and to include in Laboratory Report 1 (due two weeks form today!). Media Compositions See Chapter 3. This laboratory exercise was adapted and modified from Exercises in Cell Biology for the Undergraduate Laboratory, a project of the American Society for Cell Biology Education Committee, 1992. (http://www.ascb.org/publications/exercises.html) References Ahern, H. 1997. Introduction to Experimental Cell Biology. McGraw-Hill Companies, Inc., Dubuque, Iowa. Crupper, S.S., and J. T. Dawson. 1999. A novel approach to the isolation of plant mitochondria in the introductory college biology laboratory. Bioscene 25: 7-10.

30

Lambowitz, AM. 1979. Preparation and analysis of mitochondrial ribosomes. Methods Enzymol. 59:421-33. Zhao, J., T. Onduka, J .Y. Kinoshita, M. Honsho, T. Kinoshita, K. Shimazaki, and A. Ito. 2003. Dual subcellular distribution of cytochrome b5 in plant, cauliflower, cells. J Biochem (Tokyo) 133:115-21.

31

Chapter 5.

Cell Fractionation: Enzyme specific activity.

___________________________________

Objective To determine the specific activity of succinate dehydrogenase in the cauliflower fractions prepared two weeks ago using the data on SDH activity collected last week and a protein assay. Introduction In the exercises of the past two weeks, you used differential centrifugation to prepare a series of cell fractions. You then measured the activity of an enzyme that is known to be located in only one type of organelle in order to locate that organelle among the cell fractions. Though you only had two fractions, the resuspended mitochondrial pellet and the post mitochondrial supernatant, you probably realize that in practice a given organelle is usually found not in only a single fraction, but rather spread out among several. With modification and fine-tuning, it is possible to develop a procedure that will produce a fraction that, though perhaps not pure, is highly enriched in the organelle of interest. When assessing the effectiveness of a fractionation procedure, it is important to determine not only where a given organelle (i.e., marker enzyme) is located among the various fractions (as you have already done), but also in which fraction the majority of marker activity occurs, and also the degree of enrichment of each fraction with a given marker enzyme. Fractions will differ in volume as well as in the concentration of marker enzymes. Fractions can exhibit different enzyme activities per unit volume both because they have different enzyme content and enzyme concentration. For example, a fraction with very low marker enzyme activity might nonetheless contain the majority of the organelle if the volume of the fraction is large and the enzyme is therefore very dilute. In this case one could "correct" for the dilution factor of each fraction by calculating the total activity in the entire fraction. A hypothetical example illustrates this point: In fixedtime assays of samples from Fraction 2 and Fraction 3, both fractions yield activities of 100 g product per L of fraction. The total volume of Fraction 2 is 1.0 mL while that of Fraction 3 is 5.0 mL. Thus, the total activity of this particular enzyme in Fraction 1 is 100 mg product and in Fraction 3 it is 500 mg product. Though the assays showed the same activity per unit volume, Fraction 3 possess five times the total amount of this enzyme, and therefore of the organelle that this marker identifies. For many experiments and applications, a fraction that has both high content of a specific organelle and low contamination by other organelles is desired. For example, a fraction containing most of the total activity of a mitochondrial enzyme would be unsuitable for further experimentation (e.g., protein fractionation, electron microscopy, RNA analysis, etc....) if it also contained high levels of lysosomal enzymes. For many purposes, it would be far better to use a fraction with highly purified mitochondria, even if most of the mitochondria were found in other fractions. The purity of a marker enzyme is expressed by its enrichment. In assessing enrichment one "corrects" enzyme activity

32

for the total amount of protein present in a manner similar to that in which the fractions were "corrected" for volume in the example above. The ratio of enzyme activity to total protein is called the enzyme's "specific activity". If the activity of an enzyme is high but there is contamination by many other organelles, those other organelles will contribute protein to the fraction and the specific activity of the enzyme under consideration will be low (much enzyme activity / much protein). In contrast, a fraction with high enzyme activity and low total protein content will have high specific activity (much enzyme activity / little protein). Fractions with high specific activity are usually desired. In today's lab you will measure the protein content of your cauliflower postmitochondrial and mitochondrial fractions and then use that information, along with the SDH activity data from last week, to calculate the specific activity and total activity of each fraction.

33

A. Preparation of Protein Standard Curve. 1. Obtain 5 13 x 100 mm tubes and a tube rack. Label the tubes 1 through 5. 2. Set the spectrophotometer to 595 nm. 3. To each tube add the indicated volumes of the solutions listed in Table 1, then cover each tube with parafilm and mix by inverting slowly several times. 4. Calculate the concentration of protein in each of tubes 1-5 and write this in the space provided. The stock protein standard is 2.0 mg/mL.

Table 1. Reagent volumes used to produce a protein standard curve. The protein standard is 2.0 mg/mL. concentration Absorbance595 Tube water protein std dH2O (mLs) number (L) (L) (2 mg/mL) 1 100 0 4.0 _____________ _____________ 2 95 5 4.0 _____________ _____________ 3 90 10 4.0 _____________ _____________ 4 80 20 4.0 _____________ _____________ 5 65 35 4.0 _____________ _____________ 5. Measure the absorbance of each tube at 595 nm. 6. Save the blank (tube 1) for use in Part B below. 7. Use Excel to plot a protein standard curve. Plot a linear regression and allow Excel to calculate the equation for this line. B. Determination of Protein Content of Cell Fractions. 1. Obtain 6 13 x 100 mm tubes and a tube rack. Label the tubes. 2. Set the spectrophotometer to 595 nm. 3. To each tube add the indicated volumes of the solutions listed in Table 2 below. Cover each tube with parafilm and mix by inverting slowly several times. 4. Insert each tube into the spectrophotometer (after wiping with a kimwipe to remove fingerprints) and measure the absorbance of the mixture at 595 nm. Record your data. 5. Convert absorbances to g protein. Place this value in column G of Table 3.

34

Table 2. Reagent volumes used to determine the protein concentration of each cell fraction. Tube Fraction Amt of water Protein Absorbance595 number Fraction Reagent (L) (mL) (L) 1 Blank from Part C ____________ 2 mito. 10 90 4.0 ____________ 3 mito. 25 75 4.0 ____________ 4 mito. 50 50 4.0 ____________ 5 super. 10 90 4.0 ____________ 6 super. 25 75 4.0 ____________ 7 super. 50 50 4.0 ____________ 6. Use the data you collected last week and in this section to complete Table 3 below. Calculation of Total Activity and Specific Activity 1. Use the data collected above to complete Table 3 on the next page, and answer the questions found there.

35

Table 3. Worksheet for Enzyme Specific Activities Exercise. A B

(from Chap. 2)

C

(from Chap. 3)

D

(how much did you use?)

E

C/D

F

E·B

G

(from part B)

H

(how much did you use?)

I

G/H

J

K

J·B

L

E/J

M

=F

ENZYME tot vol of frac (mL) Mito. activity activity in the (moles/ entire fraction min · mL) (moles/min)

PROTEIN [protein] (g/L) (g/L) average protein in entire frac (mg) specific activity (moles/ total activity

Frac

slope (moles/min)

vol assyd (mL)

protein vol assyd (g) (L)

min · mg pro) (moles/min)

Super.

Questions 1. Which fraction has the most mitochondria? 2. Which fraction has the purest mitochondria? 3. Write a brief paragraph explaining the concept of "specific activity" and explain the various results one might expect

36

Chapter 6.

Enzyme Kinetics.

______________________________________

Objective To establish a saturation curve for yeast alcohol dehydrogenase, to plot the data making up the curve using three different methods, and to estimate KM and Vmax from those plots. Introduction The vast majority of reactions occurring in cells are catalyzed by enzymes. The number of different enzymes at work at any given time is very large, as are the number of substrates consumed and products produced. Yet the fundamental mechanism of catalysis is similar for all enzymes. Enzymes do not alter the equilibria between substrates and products, nor do they alter G°, they simply make the reactions go much faster than they normally would. Sometimes by a factor of a million or even a billion. Enzymes accomplish this by first binding to the substrate in a highly specific manner and then by altering the substrate molecule in such a way as to favor the formation of, and then stabilize, the transition state. The transition state is usually a unstable intermediate that can quickly break down. The spontaneous break down of the transition state molecule can lead to the reformation of the original molecule, but it can also lead to the formation of a new molecule, the product. The ability to form and stabilize the transition state is the key to enzyme catalysis. It leads to a lower activation energy of the conversion from substrate to product, and this causes the overall reaction rate to increase dramatically. There are a number of ways to study the properties of enzymes and enzyme substrate complexes. Biochemical and biophysical approaches tend to concentrate on the amino acids of the protein, particularly those that make up the active site and those that interact with cofactors and prosthetic groups. Another approach, called enzyme kinetics, focuses on the reactions themselves. These methods involve measurements of the rate of the reaction under different experimental and natural conditions. In this experiment, you will use kinetic techniques to explore the properties of alcohol dehydrogenase. Alcohol dehydrogenase The alcohol dehydrogenases (ADHs) are a family of soluble enzymes that catalyze the interconversion of simple alcohols and aldehydes using NAD+/NADH as a cofactor (figure 1). They also allow humans to consume ethanol, a highly toxic compound. The ethanol is detoxified first by conversion to acetaldehyde by ADH and then to acetic acid which is converted to acetylCoA and metabolized through the tricarboxylic acid cycle (TCA cycle, or Kreb's cycle). ADHs did not evolve so that humans could consume ethanol, but their "real" physiological role is unclear. They may have evolved in response to the ethanol created by enteric bacteria. The major vertebrate form of ADH is concentrated in the liver. Since vitamin A is an excellent substrate, it has been suggested that this vitamin is the true substrate. The aldehyde form of vitamin A is critically important in vision and for the health of some epithelial cells. ADHs are relatively nonspecific enzymes and a number of different alcohol's (e.g., n-propanol, n-butanol, npentanol) can be used as substrates.

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NAD+

NADH

O CH3CH

acetaldehyde

CH3CH2OH

ethanol NAD+ NADH

Figure 1. The interconversion of ethanol and acetaldehyde is catalyzed by ADH. Yeast (the source of the enzyme for today's lab) use ADH to produce ethanol. In these cells, the reaction of Fig. 1 proceeds primarily in the direction of reduction (i.e., to the left). The pyruvate generated from glycolysis is decarboxylated to give acetaldehyde which is then reduced to ethanol. This regenerates (i.e., reoxidizes) NAD+ and allows another round of glycolysis to occur. Assay for ADH The activity of alcohol dehydrogenases can be easily assayed by taking advantage of the fact that the oxidized form of the cofactor (NAD+) does not absorb light of 340 nm wavelength, but the reduced form (NADH) dramatically absorbs light of this wavelength. The reaction rate can thus be measured by mixing the enzyme with NAD+ and a suitable alcohol (e.g., ethanol) and then following absorbance at 340nm in a spectrophotometer. Adding ethanol as a substrate ensures that the reaction proceeds in the direction of NADH production, and A340 increases. The rate of the reaction is expressed as a change in absorbance per unit time (A340/min). By knowing the molar extinction coefficient for NADH, the actual amount of substrate oxidized can then be determined. Enzyme kinetics. Consider a simple cellular reaction in which A is converted to B by enzyme E.

A

E

B

The rate of this reaction can be measured by determining the rate of formation of B over time. The kinetic characteristics of the enzyme can be assessed by setting up a series of tubes and adding to each of them a different concentration of substrate A while adding the same amount of enzyme to each, thus holding the enzyme concentration constant (Figure 2). In each tube, the initial linear rate of the reaction is called the initial velocity and is abbreviated v0. The value of each v0 is determined as the slope of the line for each reaction; that is, the slope of each line shown in Figure 2. As the concentration of substrate (in this case A) is increased, the v0 of the reactions increase (Figure 2). If these individual reaction rates are then plotted together on a graph of reaction velocity as a function of substrate concentration, a curve like that shown in Figure 3 is produced. In most cases, the amount of product increases linearly as substrate concentration increases up to a point where the curves begins to level out and then finally plateaus. The maximum catalytic rate of the enzyme, i.e., the point at which the enzyme can work no faster no matter how much substrate is present, is call the maxim velocity and is abbreviate Vmax.

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trial #1

trial #2

trial #3

trial #4

Formation of B (or amount of B)

[A]0

[A]1 [A]2 [A]3

time

time

time

time

Figure 2. The rate of the reaction AB can be measured by measuring the rate of formation of B. The kinetics of the reaction can be assessed by holding the concentration of E constant and varying the substrate concentration of substrate, in this case [A] . The slope of each line (i.e., amount of B / unit time) is v0. One of the simplest kinetic models used to explain these phenomena is the one described in 1913 by Leonor Michaelis and Maud Menten using principles established in 1903 by Victor Henri. In the Michalis-Mento-Henri model, it is assumed that the enzyme and substrate briefly unite to form an enzyme-substrate complex that then breaks down to form product and the original enzyme. Schematically, this can be described as

. The rates of formation and breakdown of ES are described by the rate constants k1, k2, and k3. There is no "k4" because the enzyme does not recognize the product, only the substrate. According to M-M-H, Vmax is reached when sufficient substrate is present to maintain all the enzyme in the form of ES. From this scheme, and by making certain assumptions about the rate constants, Michaelis and Menten derived an expression for defining the initial velocity (v0) of an enzyme catalyzed reaction that can be used for all enzymes:

k3 k1 E + S ES E + P k2

eq. 1

Vo =

Vmax 1 + Km/[S]

=

Vmax [S] ____ Km + [S]

eq. 2

The important terms in this expression are Vmax and Km, which are unique to each specific enzyme-substrate pair. Today we call Km the Michaelis constant. It is equal to (k2 + k3)/ k1. Notice from equation 1 above that (k2 + k3)/ k1 is equal to (rate of breakdown of ES)/(rate of formation of ES). Km is a measure of the relative stability of the enzyme-substrate complex. Km and Vmax are kinetic variables. As described above, they can be determined by measuring the rate of the reaction using a fixed amount of enzyme and varying amounts of substrate. Note that Km is a concentration, not a reaction rate. It therefore has units of moles/liter, just as does substrate concentration. When v0 is equal to 1/2 Vmax, Km = [S]. Therefore Km can be determined graphically by simply determining the substrate concentration giving half-maximal

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velocity. Note that enzymes with a small Km for a particular substrate will reach saturation at a low substrate concentration. A large Km indicates that saturation is reached only at high substrate concentrations. The Michaelis-Menten equation shown above (equation 2) can be linearly transformed. That is, it can be mathematically manipulated so that it produces a straight line instead of a curve like that of Figure 3. This allows Km and Vmax to be more easily determined by using linear regression techniques. Linear plots also require fewer points (i.e., trials) to construct. One of the most common linear transformations is the Lineweaver-Burke plot. The Michaelis-Menten equation can be arranged in the form: 1/v0 = 1/Vmax + (1/[S]) Km/Vmax . eq. 3 Notice that this equation has the general form y = mx+b. A plot of 1/v0 against 1/[S] yields a straight line with a y-intercept of 1/Vmax. The intercept of the line on the x-axis is equal to -1/Km. Another useful transformation is the Eadie-Hofstee plot. In this graph, v/[S] is plotted as a function of v. The resulting line has a slope equal to -1/Km, a y-intercept equal to Vmax/Km, and a x-intercept equal to Vmax. These plots are discussed in your textbook on pages 143-145.

Vmax

Reaction velocity v0

½ Vmax

KM

substrate concentration [S]

Figure 3. A plot of the initial reaction rates (v0) of an enzyme-catalyzed reaction as a function of substrate concentration [S] yields a characteristic curve.

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Procedure Part A. Establish an appropriate ADH concentration. 1. Turn on the spectrophotometer and set the wavelength to 340 nm. 2. Dilute the ADH 1/20 with distilled water (check, this may have already been done for you). 3. Prepare a blank and an experimental tube according to table 1. Cover the tubes with a small piece of parafilm and mix by inverting several times. DO NOT ADD ETHANOL TO THE EXPERIMENTAL TUBE YET. Table 1. Preparation of tubes for establishing an appropriate concentration of ADH. Tube Blank Experimental Distilled water (mL) 3.6 3.4 pyrophosphate buffer (mL) 2.0 2.0 NAD+ (mL) 0.2 0.2 enzyme (mL) 0.2 0.2 3.0 M EtOH (mL) none 0.2

4. Zero the spectrophotometer using the blank. Save this blank for use in Part B below. 5. Start the reaction in the experimental tube by adding 0.2 mL of 3.0 M ethanol. Cover with a piece of parafilm and invert to mix. 6. Quickly insert the tube into the instrument and read and record the A340. 7. Read and record the A340 every five seconds for 30 seconds. 8. Calculate the change in absorbance per minute (A340/min) using Excel to generate a trend line trough your data. (Remember to convert from seconds to minutes.) 9. If the A340/min is between 0.1 and 0.2, continue to part B. If not, adjust the volume of enzyme (remember to keep the total assay volume equal to 6 mL). If the number is too low add more enzyme, if it is too high add less. Part B. Kinetics of ADH 1. Set up six tubes according to Table 2. Tube #1 is your blank from Part A. DO NOT ADD ETHANOL YET. 2. To determine the volume of ethanol to add to each tube use the formula Iv Ic = Fv Fc. In this case (volume needed) × (concentration of stock EtOH) = (total assay volume) × (final concentration of EtOH in the assay tube) 3. Before starting each assay, zero the instrument using tube #1 (i.e., the blank from Pat A).

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4. Perform each assay as in Part A, assaying each tube sequentially. Add ethanol to start each reaction, mix, and read every 5 seconds for 30 seconds. Repeat for each tube. 5. Convert all A340/min values to reaction velocities (µmol NAD+ reduced/min) using the following equation: total 1 A340 dilution × × assay × enzyme min factor* volume volume V0 (moles/min) =

6.22

* The dilution factor is the factor that corrects for any dilutions you made. The dilution factor can be calculated as (the volume you end up with) / (the volume you started with). For example, if you mix together 1 mL of enzyme and 1 mL of buffer, the enzyme activity (moles/min) you obtain will be one half that in the original solution because the enzyme has been diluted in half. In this case the dilution factor is 2/1 = 2, and the measured activity must be multiplied by two.

Table 2. Preparation of tubes for establishing the kinetics of ADH. Tube [ethanol] # (M) 1 2 3 4 5 6 7 0.000 0.025 0.050 0.100 0.150 0.200 0.300 3M stock ethanol* 0.000 _______ _______ _______ _______ _______ _______ Pyrophos. buffer (mL) NAD+ (mL) enzyme (mL)** Part A _______ _______ _______ _______ _______ _______ Distilled water *** here. _______ _______ _______ _______ _______ _______ Total Vol. (mL) = 6.0 = 6.0 = 6.0 = 6.0 = 6.0 = 6.0 = 6.0

Blank from 2.0 2.0 2.0 2.0 2.0 2.0 _______ _______ _______ _______ _______ _______

* Determined using the equation in step B2 above. ** Determined from Part A above. *** Add sufficient water to bring the final total up to 6.0 mL.

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6. Plot your results as 1) a saturation curve (like figure 3 above), 2) a Lineweaver-Burke plot, and 3) an Eadie-Hofstee plot (see pages 143-145 in you textbook). NOTE: Do not include the 0,0 points in the linear transformation plots. 7. Estimate or calculate Km and Vmax from your plots. 8. Prepare a table comparing the values of Km and Vmax obtained from the three plots.

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Chapter 7.

Protein Fractionation: Purification of IgG from Human Serum.

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Objective To isolate immunoglobin G from human serum using ion exchange chromatography. Introduction In a previous exercise, you fractionated cells into their component organelles. In this exercise you will perform a fractionation at the molecular level. Here, you will isolate a single protein from a mixture of scores of other proteins. Obviously, this can be very useful. For example, you might wish to examine the kinetics of a particular mitochondrial enzyme (like succinate dehydrogenase), or determine the amino acid sequence of a G-protein. There are any number of reasons why you might need to isolate a single protein. Once you have isolated the protein (or, more precisely, believe you have isolated the protein) it is important that you verify your results. That is, you must be able to produce evidence that 1) you have isolated a single protein form the mixture of hundreds, or not, and 2) that the single protein you have isolated is the one you think it is. There are a large number of methods for purifying proteins. Typically, proteins are purified from the cell fraction that would contain the greatest amount of the protein. For example, DNA or RNA polymerases would be isolated from the nuclei, glycolytic enzymes from the cytoplasm, enzymes of the tricarboxylic acid cycle from mitochondria, etc. Therefore, before protein fractionation, several steps of cell fractionation are usually involved. Since you have already performed a cell fractionation procedure, we will perform the protein fractionation using a relatively simple starting material. In this exercise, the protein that you will isolate will come not from cells but from serum, that is, the fraction of blood that remains liquid after the blood has clotted. The protein you seek is an immunoglobin (also called an antibody) named immunoglobin G (IgG). Immunoglobins are produced by cells of the immune system in response to foreign material. They are a major component of serum, second only to serum albumin in abundance. They function by binding specifically to heavy chains foreign proteins and bacteria, resulting (usually indirectly) in their inactivation and destruction. IgG is one of five classes of immunoglobins: IgG, IgM, IgA, IgD, and IgE. Each class of immunoglobin is light chains characterized by specific functions and structural features. IgG provides resistance against many viruses, bacteria and bacterial toxins. IgE accelerates local inflammation disulfide bonds and is responsible for allergic reactions in people prone to them. IgD is found on the surface of a type of immune cell called B cells. IgM is the first type of immunoglobin Figure 1. The structure of IgG.

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secreted after an antigen is detected. Finally, IgA is less well understood, and is primarily found in glandular secretions. Structurally, each immunoglobin class has a unique composition of heavy chain and light chain isotypes. In some species, the immunoglobulin classes are further differentiated into subclasses. For examples, in humans, IgG antibodies comprise four IgG subclasses, IgG1, IgG2, IgG3, and IgG4, and there are two IgA subclasses, IgA1 and IgA2. In many antibodies, including the IgGs, the heavy and light chains are held together by disulfide bonds. The structure of IgG is shown in Figure 1. To learn more about immunoglobins and the immune system, take Biol 333, Immunology. Protein Fractionation Regardless of the starting material (i.e., cell culture, serum, etc...) and the initial processing steps, ultimately the desired protein is obtained in some type of buffer solution together with a large number of unrelated proteins. At this point, the protein can be subjected to a number of techniques that can, in principle at least, separate it from all other proteins in the sample. There are many such techniques available in the literature. If the protein you wish to isolate has never been isolated before, your choice of which method(s) to use will depend on many variables and might involve a considerable amount of empirical testing. Some of the more widespread methods are described below. Fractionation by precipitation. It is possible to alter the composition of the buffer to force the precipitation of a portion of the proteins in solution. This method takes advantage of the differential solubility of proteins under different solute compositions. One popular method is called "salting out" and involves adding increasing amounts of ammonium sulfate to the protein solution. As the ionic strength of the solution rises, proteins and salt compete for the residual water, and proteins begin to interact via hydrophobic patches on their surfaces which causes them to precipitate. Ammonium sulfate is used as the salt of choice because it preserves protein activity and promotes precipitation at lower concentrations than other salts. Another precipitation method is isoelectric precipitation. The solubility of proteins depends not only on ionic strength, but also on pH. Recall that the charge state of proteins (in reality the charge state of the amino acid side groups) is pH dependent. At low pH, proteins become more protonated, which tends to make them more positively charged, while at high pH the side H2N - CH2 - COOH3N+ - CH2 - COOH groups are more depH=12.0 pH=1.0 protonated and thus the protein as a whole tends to become more negatively charged. At one particular H3N+ - CH2 - COOpH, the protein carries a net charge of 0 as the number of pH=7.0 negative charges equals the Figure 2. Amino acids can be ionized number of positive charges. This pH is called the isoelectric point (pI) and its value is unique for each protein. Most proteins are highly soluble at physiological ionic strength and pH (in some tissues, soluble proteins make up 40% of total cell mass). If the pH of the experimental buffer is altered to approach the pI of a given protein, the

+

-

0

43

net charge on that protein will go to zero, and the absence of electrostatic repulsion from other proteins can lead to protein aggregation and precipitation as weak electrostatic attractive forces, such as van der Waal's interactions, prevail. The tendency of proteins to interact near their pIs can be promoted by reducing the ionic strength or by adding organic solvent. Thus the technique of solvent precipitation can be useful. In this technique, organic solvent (usually ethanol or acetone) is added to the solution of proteins in order to reduce water activity. This reduces the water available for protein solvation and causes protein aggregation similar to that seen in salting out and isoelectric precipitation. Fractionation by dialysis and ultrafiltration. Semipermeable membranes can be used to separate proteins based on their size. In dialysis, the protein sample is enclosed in a tube of semipermeable membrane and exposed to a large volume of a desired buffer. The low-molecular weight compounds (buffering agents, salts, etc...) pass freely through the membrane pores whereas the protein is retained. Ultrafiltration is similar except that the pores can be larger and therefore allow smaller proteins to pass through. Increasingly large pore size (molecular weigh cutoff) allow increasing large proteins to permeate. Chromatographic and electrophoretic fractionation procedures. Chromatography refers to any method that solutes are fractionated by partitioning between a mobile phase (a solvent or buffer) and an immobile phase (the matrix). Among these techniques are gel filtration and adsorption chromatography. Gel filtration (also called size exclusion or gel permeation chromatography) separates molecules (including proteins) on the basis of their size and shape. In this case the matrix is made of beads of a material containing many pores of different sizes (think of Swiss cheese or a kitchen sponge). As the protein solution is allowed to flow over the matrix beads, large proteins are unable to enter the matrix, and must flow around the beads in the interstitial spaces between them. This space is known as the void volume, V0. For most gel filtration resins, V0 is about 1/3 of VT, the total volume of the column in which the matrix is packed. These larger molecules, therefore, travel only through V0 before eluting at the end of the column. Molecules smaller than the pore size diffuse into the matrix beads and must travel the entire volume VT before eluting. In other words, large proteins confined to the interstitial space will migrate more rapidly through the column than small molecules which diffuse into and back out of the matrix and consequently are partly trapped and fall behind. Since the pores vary in size, proteins of intermediate size will penetrate to varying degrees into the beads and thus are separated from each other on the basis of their size. Proteins of equal mass but different shapes will differ in their apparent size. For example, an elongated protein will appear larger. It will have a larger "tumbling radius" and will move through the column more rapidly than a spherical protein of the same mass. In adsorption chromatography, the molecules of interest actually bind to the matrix material (this is called adsorption), and differences in the adsorption affinities of the molecules are used to separate them. In general, the adsorption behavior of a specific protein can be described by a partition coefficient . The value of represents the fraction of the protein that is adsorbed. It can therefore have values between 0 (no binding) and 1.0 (complete adsorption). An aqueous solution of proteins (the mobile phase) is allowed to pass over the adsorption matrix (the matrix, or stationary phase). The proteins then preferentially interact with each phase. Some bind tightly to the matrix, others do not bind at all and flow with the buffer front. Still other protein (the majority in most cases) exhibit some intermediate affinity for the matrix, and their progress through the column is hindered to some extent. One type of adsorption chromatography, the one that you are going to use to isolate IgG,

44

is ion exchange chromatography. In this type of chromatography, the charged amino acid side groups on the surfaces of proteins bind to an insoluble matrix with the opposite charge. More precisely, the ionized proteins displace the counter ions (e.g., chloride or sodium) electrostatically bound to the functional group of the matrix. In turn, the proteins can be displaced by increasing the concentration of ions in the elution buffer. A highly effective method involves using changes in pH, and therefore using protons (H+) as the counter ion. Since the overall charge of a protein is a dependent on pH (see above), a pH gradient can be employed so that the net charge on the adsorbed protein changes. Under specific starting conditions of buffer, pH, and ionic strength, the net charge on the protein of interest can be manipulated to interact with the matrix. In a mixture of proteins, the total tissue (or cell, or organelle) extract can be loaded onto the column at low (or high) pH in a low (or high) pH buffer so that all the protein bind strongly to the matrix. As the pH of the eluting buffer is increased (or decreased), proteins with different pIs will begin to elute. Theoretically, each protein will elute at its characteristic pI and complete resolution of all proteins can be achieved. In reality, this is rarely the case because many pIs are close to one another, and the proteins do not all suddenly detach at the same moment and at exactly at the same pH. Thus, the most important parameters to consider in an ion exchange separation are the choice of ion exchange matrix and the initial conditions of pH and ionic strength. It has been estimated that up to 80% of proteins are negatively charged at neutral pH. Consequently, the most used ionic adsorbent matrices are those with functional groups that are positively charged at neutral pH. We use these facts to our advantage in this exercise, but in the "reverse" sense. The proteins of human serum are all negatively charged at neutral pH. All, that is, except IgG. IgG is positively charged at neutral pH. Therefore, IgG can be very effectively isolated by passing a neutral solution (pH = 7.0) of serum proteins over a column containing a positively charged matrix. When the column is eluted with a buffer of pH 7.0, all the (negatively charged) serum proteins bind to the (positively charged) matrix except IgG, which flows out with the buffer front. In this type of separation, it is not necessary to actually do the separation in a column. It is much easier, and equally effective, to simply mix the protein solution and the binding matrix and allow binding to occur in a test tube. This is called batch processing or batch mode. The ion exchange matrix can be loaded in batch mode by mixing the matrix and the sample in a tube and then stirring or shaking the mixture. Subsequently, the matrix is poured into a column for gradient elution, or batch eluted on a sintered glass funnel. Alternatively, as is the case in your separation, the buffer containing the IgG can simply be decanted into a new tube, and the matrix containing the unwanted proteins can be discarded, or washed and reused. Protein Fractionation in BIOL 350 is a three week process This exercise consists of three parts (one per week). In the first week you will isolate IgG by ion exchange chromatography and determine the protein concentration of the resulting protein preparation. In the second week you will separate the proteins by sodium dodecyl sulfate polyacrylamide electrophoresis (SDS-PAGE) and determine how many different proteins are in your protein preparation and their sizes (molecular weights). In the third week you will unequivocally determine whether your protein preparation contains IgG or not using Western blotting techniques. All of these techniques are used routinely in Cell Biology laboratories around the world, including the one that might hire you after graduation. So make a concerted

45

effort to understand these techniques well so you can show off your knowledge on your resume and at your job interview!

46

Precautions The human serum you will be using was purchased from a company that has tested it to be certain it is free of hepatitis B and C, and HIV, however, you should still treat it as you would any human blood product. For safety reasons, consider it potentially infectious. Wear gloves, and dispose of items that touch the serum in the bleach container provided or in the orange biohazard bag.

Procedure Part A. Dialysis of human serum. 1. Dialyze 30 mL of human serum against 0.01 M phosphate buffer (pH 7.2) for at least 24 hours at 4°C with constant stirring. Change the entire buffer volume at least once, preferably at 9-12 hours. Each group will require 2 mL of dialyzed serum. Part B. Preparation of DEAE cellulaose 1. To prepare the diethylaminoethyl (DEAE) cellulose, follow the instructions in the Whatman DEAE manual precisely.

Part C. Protein fractionation by bulk phase ion exchange chromatography. 1. Add 1 mL of dialyzed human serum to 10 mL of DEAE cellulose in a 15 mL centrifuge tube. Mix the contents of the tube by inverting it several times. 2. Incubate the bulk phase IEC matrix for 1 hr at room temperature. Mix by repeated inversion every 5 minutes. 3. After the incubation period, centrifuge the tube at 1,000 g (2300 rpm on the Baxter Megafuge) for 10 minutes. 4. Immediately after centrifugation, use a transfer pipette to place the supernatant into clean 15 mL culture tube. Measure the volume of your sample. (Why do you need this information?) The DEAE (with the proteins bound to it) can be discarded. 5. Transfer approximately 0.5 mL of the supernatant to a microfuge tube and set aside. Label the tube containing the remainder of the supernatant and place it in the freezer (-20°C) until next week. Part D. Determine the protein concentration of your putative IgG sample. D1. Preparation of a standard curve. 1. Obtain a BSA standard that is 2 mg/mL. 2. Turn on the spectrophotometer and set the wavelength to 595 nm. 3. Obtain six 13 x 100 mm test tubes and label them #1-6. 4. Prepare a series of standards in the six tubes according to Table 1.

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Table 1. Reagent mixture for preparation of a protein standard curve. The BSA atandard is 2 mg/mL. BSA std (µL) Amount of BSA Tube H2O (µL) (mg) 1 100 0 _________ 2 95 5 _________ 3 90 10 _________ 4 80 20 _________ 5 65 35 _________ 6 50 50 _________

5. Add 4.0 mL of BioRad protein assay reagent to each tube. Cover each tube with parafilm and mix by inversion. 6. Insert tube #1 into the instrument and set zero. (Tube #1 is the "blank", it contains 0 mg protein. You are setting the instrument to read 0 absorbance when there is 0 mg protein.) 7. Read the absorbance at 595 nm (A595) for tubes #2-6. 8. Using the Excel program on the computer at your work station: a. Plot your standard curve. b. Have the program calculate the equation for the line that describes your data D2. Determine the protein concentration of your sample and the original serum sample. 1. Label eight 13 x 100 mm test tubes. 2. Prepare dilutions of your sample (in the microfuge tube) as described in Table 2 below. Table 2. Reagent mixture for assaying protein content of IgG samples. IgG sample original serum Tube H2O (µL) (µL) (µL) 1 100 0 --2 90 10 --3 75 25 --4 50 50 --5 25 75 --6 0 100 --7 95 --5 8 90 --10 3. Add 5.0 mL of BioRad protein assay reagent to each tube. Cover each tube with parafilm and mix by inversion.

48

4. Insert tube #1 into the instrument and set zero. (Again, tube #1 is the "blank" because it contains 0 mg IgG.) 5. Read the absorbance at 595 nm (A595) for tubes #2-8. 6. Using your standard curve, determine the protein concentration of tubes #1-8. Don't forget to convert to concentrations! Questions. 1. Should the A595 values of tubes #1-6 in part D2 above have been the same? (After all, they all contained just water and IgG solution.) 2. Should the protein content of each sample placed in tubes #1-6 have been the same? 3. Should the protein concentration of each sample placed in tubes #1-6 have been the same? 4. Based on your answer to question #3, why did you prepare six tubes of the IgG solution. I.e., in Part D2 above, why didn't you just prepare one of these? 5. What is the protein concentration of the original serum? (Remember, concentration is different from content.) What is the total protein content of the original serum? 6. What is the protein concentration of your putative IgG sample? What is its total protein content? 7. Did the DEAE cellulose matrix remove any protein from the serum? How much?

References Reisfeld, R.A. et al. 1962. Acidic buffer system for resolution of cationic proteins. Nature 195:281.

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Chapter 8.

Protein Fractionation: Sodium Dodecyl Sulfate ­ Polyacrylamide Gel Electrophoresis.

___________________________________

Objective To separate proteins based on their molecular weight using SDS-PAGE. Introduction Last week you purified IgG from human serum by ion exchange chromatography and determined the amount of protein in your resulting sample. You also determined the protein concentration of the original serum, and were asked to determine if the ion exchange procedure had removed any proteins from the sample. You now have a tube containing an aqueous solution of IgGs. At least that's what you think you have! How do you determine what is actually in your sample? One possible first step is to carry out a procedure called sodium dodecyl sulfate ­ polyacrylamide gel electrophoresis, or SDS-PAGE. This procedure is described on page 180 of your text. There are also a large number of web sites that describe and explain this procedure. In essence, SDS-PAGE allows you to determine the molecular weights of the all the proteins in your sample. You can then compare the molecular weight of your target protein (in this case IgG) to the molecular weights of the proteins present in your sample to determine whether or not the procedure has been successful. As the name indicates, SDS-PAGE uses polyacrylamide. When acrylamide is induced to polymerize, it forms a gel with a mesh-like structure. This polyacrylamide gel acts as a molecular sieve. Small proteins are able to readily enter the mesh and move rapidly though the gel. Larger proteins are hindered and take longer to pass through. Thus SDS-PAGE separates proteins according to their molecular weight. Unlike chromatography, however, the molecules are forced to pass through the gel not by the flow of a mobile phase, but by their electrostatic attraction to a electrode. The first step in the preparation of a sample for SDS-PAGE is to denature all its proteins in the presence of an anionic detergent. This step gives the proteins a uniform negative charge, and denaturation assures that movement of the protein is independent of its original shape. Denaturation is achieved by boiling, and the anionic detergent that is almost universally included in this step is sodium dodecyl sulfate (SDS). The compound -mercaptoethanol (-ME) is also added here to ensure the complete reduction (i.e., breakage) of any disulfide bonds. The proteins that emerge from this step, and that are loaded onto the polyacrylamide gel, are essentially linear and covered with negative charges. The gel itself is not prepared in a tube as in column chromatography, but rather it is formed as a thin sheet between two layers of glass. The protein sample is loaded onto the edge of this "slab" gel. Once loaded onto the gel, a positive electrode is placed at the opposite edge and the negatively charged proteins migrate towards it, through the gel between the two sheets of glass, due to electrostatic attraction. The gel system that you will be using is a "discontinuous" gel. In this type of gel, the proteins first enter a "stacking gel" that is prepared to have very low porosity (very large mesh size). This initial gel causes the proteins to "stack" or concentrate into a single band at the top of

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the resolving gel. When they reach the resolving gel, they then separate according to molecular weight. The proteins moving through the gel can not be seen, so before being loaded onto the stacking gel, a blue dye is added so the progress of the dye front through the gel can be visualized. Once the proteins have been separated, they also need to be visualized, which is accomplished by staining. There are a number of different stains, but Coomassie Blue is by far the most popular and frequently used. The staining solution also contains a fixative that immobilizes the protein bands within the gel. The stain is also absorbed by the gel, so the gel must be "destained". Once the gel is stained and then destained, the results of the separation can be documented by digitally photographing the gel on the light table and storing the image on disk. Procedure Apparatus Electrophoresis chambers Power supplies Micropipettors Water bath set to boiling (or a beaker in the microwave) Digital camera/light table/computer set up Supplies IgG sample from last week Sample buffer (1% bromphenol blue/SDS/glycerol/-ME) Running buffer (10x tris/glucine/SDS from Bio-Rad) Coomassie blue stain (Bio-Safe Coomassie for Bio-Rad) Microcentrifuge tubes Gel loading tips A. SDS-PAGE. 1. You are going to prepare two identical gels, one to be used for Coomassie staining and the other for next week's Western blotting exercise. Two gels (one electrophoresis apparatus) will be shared by the four groups of each bench. It is important that each gel be identical, so be careful about how you load it (i.e., which sample goes into which well on each gel). 2. From the results of the protein assays you performed last week, calculate the volumes of your sample and of diluted serum that contain 50 g of protein. For example, if you determined that your sample contained 8 g protein/L, then set up the ratio:

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1 L 8 g 50 g 8 g x

=

x L 50 g x L

and solve for x

=

=

6.25 L

In this case, you would need to take 6.25 L of IgG solution in order to get 50 g of total protein.

3. Add the appropriate volume of your sample and of diluted serum to two separate microcentrifuge tubes. (Only 1 group needs to prepare the diluted whole serum sample.) 4. Add 1/4 volume of Sample buffer to each tube and mix well. For the example above, add 6.25/4 = 1.56 L of Sample buffer. 5. Float the tubes (in a floating rack) in the boiling water bath for 3 min. Then allow them to cool on ice for 1 min. 6. Using gel loading tips, load the samples onto the two gels (load half of each sample into each gel, but add no more than 30 L into any one well). You will be shown how to do this. Load the gels as follows: Lane 1 2 3 4 5 6 7 8 9 10 Sample no sample molecular weight standards (5L) no sample diluted whole serum no sample Group 1 IgG sample Group 2 IgG sample Group 3 IgG sample Group 4 IgG sample no sample

Repeat exactly for gel #2. 7. Add running buffer to the appropriate level, connect the power supply, and electrophorese the samples until the blue dye reaches the bottom of the gel. A good setting is 200 volts for about 45 min. 8. Once the two gel have run to completion, turn off the power and remove the gels from the apparatus. (You will be shown how to do this).

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9. Remove gel #1 from between the glass plates (wear gloves and be careful because the gel can tear!). Stain the gel in the Coomassie solution. 10. Destain gel #1 (The specifics of steps 8 and 9 will depend on the Coomassie stain we will be using [we change often to try new products]. Details will be provided before the class starts.) 11. Go to part C below using gel #2. B. Determination of the molecular weights of the proteins present. 1. Using Excel on the computer at your workstation, plot a standard curve based on the (hopefully) linear relationship between the logarithm of the molecular weights of the standards (provided to you on a card from the manufacturer) and the distance each protein migrates from the bottom of the loading well. a. For each protein in the molecular weight marker lane, measure the distance (in mm) from the top of the resolving gel to the center of each band. b. Measure the distance form the top of the resolving gel to the dye front. c. Calculate the relative mobility (Rf) of each protein standard as: Distance of protein migration Rf = Distance of dye front migration

d. Plot Rf versus log10 molecular weight. Have the software perform a linear regression, plot the regression line, and calculate the equation that describes the best fit line. e. Calculate Rf for each protein band that appears in your gel. Determine the molecular weight of each protein. f. You will also use this procedure in the next Chapter (Chapter 9) to determine the molecular weights of the proteins visible on the western blot. C. Transfer to nitrocellulose. 1. Remove gel #2 from the between the glass plates and place it in Transfer buffer for 15 min. 2. While the gel is equilibrating with the transfer buffer, build the transfer apparatus. The assembly must be done in the following order: a. Against the black backing of the clamping unit place a scotch-brite pad b. Against the scotch-brite pad place a piece of presoaked (in transfer buffer) filter paper. c. Carefully place gel #2 on top of the filter paper. d. Place the nitrocellulose membrane on top of the gel. (Do not touch the nitrocellulose without gloves.) Make sure there are no bubbles between the nitrocellulose and the gel. Gently roll a pipette over the surface to remove bubbles. e. Place another piece of presoaked filter paper on top of the nitrocellulose.

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f. Place another scotch-brite pad on top of this. g. Close and clamp the unit. h. Place the clamp in the transfer chamber. Carefully check the orientation! Make sure the proteins (negatively charged) are being pulled by the positive electrode into the nitrocellulose, otherwise they will simply vanish into the transfer medium, never to be seen again. i. Transfer the proteins onto the nitrocellulose at 30 volts overnight. Go back to part B above. Questions 1. How many proteins are present in the serum? How many are present in your IgG sample? 2. Do all of the protein bands that appear in the lane with your IgG sample also appear in the lane with whole serum? If not, how do you explain this observation? 3. Your IgG sample contains a number of bands, not just one, as you may have been expecting (one band corresponding to IgG). Does this necessarily mean that proteins other than IgG are present? Explain (Hint: What does the structure of IgG and the way you treated the sample for SDS-PAGE suggest you should find?) 4. Use the internet to find the molecular weights of IgG as a whole, and its heavy and light chains. 5. Do the molecular weights of the proteins in your IgG lane conform to a pattern of mixed heavy and light chain subunits? 6. Based on your results, is ion exchange chromatography a suitable method for isolating IgG from human serum? References Laemmli, U.K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680. Matsudiara, P.T. and D.R. Burgess. 1987. SDS microslab linear gradient polyacrylamide gel electrophoresis. Anal. Biochem. 87:386-396. Reisfeld, R.A. et al. 1962. Acidic buffer system for resolution of cationic proteins. Nature 195:281. Towbin, H. et al. 1979. Electrophoretic transfer of proteins from SDS and acid/urea gels to nitrocellulose. Proc. Natl. Acad. Sci. USA. 76:4350-4354. Weber, K. and J. Osborn. 1969. Basic study of molecular weight determination by SDS electrophoresis in a continuous buffer system. J. Biol. Chem. 224:4406-4412.

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Chapter 9.

Protein Fractionation: Western Blotting.

___________________________________

Objective To identify a protein based on immunoglobin binding. Introduction Western blotting (also called Immunoblotting) is a very powerful technique that combines the resolving power of gel electrophoresis with the detection specificity of immunoglobins. The name of the procedure comes form E. M. Southern, who developed a procedure to bind single stranded DNA to a membrane and then identify it by probing it with RNA of known sequences. This method became known as "Southern blotting", and is capitalized because it is derived from a proper name. Soon afterwards, a similar "blotting" technique was developed to identify specific RNA molecules bound to a membrane by probing them with other RNA molecules. The people who developed this technique called it "northern blotting" as a humorous twist. When someone then developed a method to identify immobilized proteins by probing them with specific immunoglobins (antibodies), it was, predictably, called western blotting. As of today, there is no method called eastern blotting. Recall that immunoglobins are proteins produced by the immune system as a defense mechanism. They recognize and bind to a particular arrangement of amino acids on a protein. They thus bind to proteins located on the surface of cells and viruses in a highly specific manner. This specificity of binding is exploited in the technique of western blotting. By injecting a particular protein into, a rabbit or a goat, or any other vertebrate, the animal's B cells will produce immunoglobins to that protein. The target of an immunoglobin is called its antigen. The immunoglobin can then be isolated from the animal's blood and used as a probe. It possesses the innate ability to will bind specifically to its antigen, and nothing else. For example, we could run a SDS-PAGE, electrostatically transfer the proteins from the gel onto nitrocellulose (which is physically stronger than the gel and will last longer; it also tightly binds all proteins), then wash the nitrocellulose with a solution containing an immunoglobin to a specific antigen to determine if that particular antigen is present on the nitrocellulose. This is precisely what you have done over the last two weeks. You think you have isolated a protein (IgG) from human serum. You have performed SDS-PAGE and have determined that the molecular weights of the proteins present in your sample are consistent with them being IgG subunits. Now you are going to unambiguously identify them by adding a human IgG specific immunoglobin. A company has injected human IgG into a goat and made, and now sells, an antibody to IgG. (Don't become confused here, you are using an immunoglobin to detect another immunoglobin. Immunoglobins can be used to detect any protein, including other immunoglobins!) There are of couple of complexities to the western blotting technique that must be addressed. First, as you observed, the proteins migrating on an acrylamide gel are not visible. Nor are they visible on the nitrocellulose (the colored standards are visible, but nothing else). You can not see the immunoglobins, nor any of the other proteins. You had to stain the gel in order to detect them. So if you add an immunoglobin to IgG, it will bind very nicely, but you can't see it! In order to visualize the IgG on the ntrocellulose, you will add a immunoglobin that

55

has been modified by biotechnology. The immunoglobin you will use has the enzyme horseradish peroxidase (HRP) conjugated to it. It has the antigen binding region intact, and on the other end is attached the enzyme. The immunoglobin is a "goat anti-human IgG HRP conjugate". After the conjugated immunoglobin is added and binds to its antigen (the human IgG), the substrate for HRP is added. The product produced by the enzyme forms a purple precipitate on the nitrocellulose, and in that way allows detection of the IgG band. The other complexity concerns the use of nitrocellulose. We chose to use nitrocellulose to immobilize the proteins from the gel because it has very strong protein binding properties. If one were to add the HRP-conjugated immunoglobin to the nitrocellulose without some type of pretreatment, it will specifically bind to its antigen, as expected, but it will also non-specifically bind to the nitrocellulose by virtue of it simply being another protein.. This non-specific binding can be blocked by first soaking the nitrocellulose in a solution of protein (you will use BSA, many labs use powdered milk for this purpose). This treatment completely covers all the nonspecific binding sites with protein. After such a pretreatment, the HRP-conjugated immunoglobin will only bind specifically to is antigen. Procedure Be sure to wear gloves during this procedure, and be careful not to touch the nitrocellulose. 1. Add 20 mL of Blocking Solution to the nitrocellulose in a incubation tray. Incubate for 60 min with rocking. 2. Discard the blocking solution down the sink and rinse the membrane for 5 min with 20 mL of Wash Buffer. 3. Discard the Wash buffer (down the sink) and add 20 mL of an appropriately diluted solution of goat anti-human IgG HRP conjugate. Incubate for 40 min with rocking. 4. Discard the immunoglobin solution (down the sink) and rinse the nitrocellulose for 5 min with 20 mL of Wash Buffer. Discard and repeat this wash step two more times. 5. After discarding the final wash solution, add 20 mL of the color development solution (the HRP substrate). Bands may appear immediately, and color development should be complete by 15 min. To stop color development, wash the membrane with several changes of distilled water. 6. Measure the distance traveled by each of the proteins made visible by the IgG HRP conjugate, calculate Rf for each, and plot Rf versus log10 molecular weight as described in Part B of Chapter 8.

Questions. 1. Do you detect IgG on your blot? Do you see bands representing both heavy and light chains? Explain how you might be able to visualize only one or the other using the western blotting technique.

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2. Do the molecular weights of the proteins on your blot correspond to those you found in the Coomassie stained gel? 3. Do you see any other bands on the gel? If you do (of if you did) how would you explain their detection? 4. Explain the following (theoretical) results: a. The gel stained with Coomassie shows a band that does not appear on the western blot. b. The western blot shows a band that does not appear on the Coomassie stained gel.

References Laemmli, U.K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680. Matsudiara, P.T. and D.R. Burgess. 1987. SDS microslab linear gradient polyacrylamide gel electrophoresis. Anal. Biochem. 87:386-396. Reisfeld, R.A. et al. 1962. Acidic buffer system for resolution of cationic proteins. Nature 195:281. Towbin, H. et al. 1979. Electrophoretic transfer of proteins from SDS and acid/urea gels to nitrocellulose. Proc. Natl. Acad. Sci. USA. 76:4350-4354. Weber, K. and J. Osborn. 1969. Basic study of molecular weight determination by SDS electrophoresis in a continuous buffer system. J. Biol. Chem. 224:4406-4412.

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Chapter 10.

Cell Culture: Determining the Density of Cell Cultures.

___________________________________

Objective To use a hemocytometer to determine the density of cells in a cell culture. Introduction One of the most important advances in the study of cell biology was the discovery that cells of metazoans, particularly higher plants and animals, could be grown in a flask, outside of the original organism. The first cells to be successfully cultured in vitro were nerve cells. In 1907, R. G. Harrison, who sought to show that nerve fibers grew from individual neurons, explanted a piece of neural tube from a frog embryo into a drop of clotted lymph on a cover slip. His observations that axons did indeed arise from individual neurons helped establish the concepts of nervous system structure that are still accepted today. His experiments also proved that cells could survive and even grow and develop in culture. One hundred years later, few techniques are as fundamental to the study of cells. Over the next four weeks, you will become familiar with some of the concepts and methodologies that are central to successfully culturing cells in the laboratory. "Tissue culture" is often a generic term that refers to both organ culture and cell culture. These terms are essentially synonymous. Cell cultures derived from organs or other primary sources, that is, from the original animal source, are called primary cultures. Secondary cultures are those that are derived from primary cultures. In general, cultures of normal cells grow and survive only through a finite number of cell divisions before senescing and dying. For animal cells, 50-100 divisions appear to be the upper limit. The loss of vigor and eventual cell death is termed replicative senescence. In contrast, some stem cells and cancerous cells proliferate indefinitely in culture. These cells are therefore called "immortal cells". Both types of culture have contributed greatly to our understanding of cell biology, In this week's laboratory exercise, you will learn one of the most fundamental of all cell culture techniques: determining the cell density of a culture using a hemocytometer. In a majority of experiments involving cells in culture, it is necessary to know how many cells are present in the experiment. Cells in culture are of two fundamental types, and the way you prepare them for counting is different. Cells are either adherent or non-adherent. As the name suggests, adherent cells normally prefer to be attached to the substratum (both in the original animal and in the culture flask). During interphase, adherent cells in culture, particularly fibroblast-like cells, spread out on the substratum in a characteristic, "spindle-shaped" configuration (Figure 1). They detach from the substratum to undergo mitosis. Non-adherent cells, which are more uncommon, do not attach to the substratum. Typically, these cells, in their original state, were circulating cells such as white blood cells. Because they are not attached, these cells are the easiest to prepare for counting. A sample can be taken after simply mixing the flask. Adherent cells require slightly more involved procedures. One of the first steps often involves the use of a calcium- and magnesium-free medium and a proteolytic enzyme to detach the cells from the culture vessel. The absence of calcium and magnesium in the wash medium

58

helps remove these ions from the adhesion disks cells use to attach to the substratum. Part of the holding ability of these disks arises from electrostatic interactions between negatively charged proteins in the disks and the negatively charged surface of the substratum. The positive charges of divalent cations act as an adhesive to hold the negative charges together. The proteolytic enzyme, usually trypsin, collagenase or pronase, helps release the cells by destroying the proteins of the adhesion disks. Of course, all other proteins of the cell surface are also digested, so these enzymes must be used with caution to avoid excessive cell damage. After the majority of cells have detached, they are washed with medium containing serum, which has natural inhibitors of proteolytic enzymes. Some supply companies include a chelating agent such as ethylenediaminetetracetic acid (EDTA) in the trypsin mixture. These are agents that bind to free calcium and magnesium and take them out of solution.

Figure 1. A fibroblast in culture. Use of the Hemocytometer Counting cells with a hemocytometer is an easy, fast, and efficient way to establish the density of a cell culture. A hemocytometer is a glass plate that looks like a regular, but very thick, microscope slide (Figure 2). On its surface are two precisely ground counting areas that can be seen best under a microscope. Each of these "counting chambers" is divided into nine 1.0 mm squares (Figure 3). When placed on top of the counting chambers, a special glass cover slip (not a normal cover slip for slides) defines an area that is exactly 0.1 mm deep. Thus the total volume of each square in the counting chamber [there are 9 of these squares (Figure 1)] is 1.0 mm × 1.0 mm × 0.1 mm, or 0.1 mm3, which is also 10-4 cm3. Since 1 cm3 is equal to 1 mL, the density of cells per mL (i.e., the concentration of cells) is equal to the number of cells in each 1 mm2 square times 104.

Figure 2. The hemocytometer.

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Though counting cells on a hemocytometer is easy, there are several sources of error that must be avoided. The most obvious is to be sure to account for any and all dilutions that were made to the original cell culture before counting. In addition, hemocytometer counts are subject to error if: 1. The cells in the original sample are not homogenously distributed. If the cells in the original sample are clustered or clumping together, accurate counts are difficult or impossible. 2. The counting chamber is improperly filled or if an inappropriate cover slip is used. The depth of the counting chambers must be precise. Over- or underfilling them, or using a cover slip that sags, will introduce significant errors to your counts. 3. Counts are taken improperly. There is a convention for counting cells that should always be adhered to. For example, if a cell is "on the line" is it counted or not? Also, the statistical vigor of your counts should not be overlooked. The details of these conventions, and ways to avoid them, are discussed below.

A

}

1 mm

Figure 3. The counting chamber of a hemocytometer is etched to contain a series of grids. The central grid, used for counting small cells, is labeled "A". The corner grids, those with 16 small squares each, are used to count larger cells.

The error between cell counts can be reduced to as little as 15% if simple precautions are taken. For example, it is assumed that the sample present in the counting chamber is a random sample that faithfully reflects the contents of the original cell culture. This assumption will be invalid if the sample is taken from a culture in which the cells are clumping or that has not been mixed well. Unless 90% or more of the cells are free from contact with other cells, the

60

count should be repeated with a new sample. Also, a sample will not be representative if the cells are allowed to settle before the sample is taken. Always mix the cell suspension thoroughly (but gently) before sampling. The cell suspension should be diluted so that each 1 mm2 square contains between 25250 cells (i.e., a beginning concentration of 2.5 ­ 25 × 105 cells/mL). For statistical reasons, more than 200 cells should be counted (the counting error is approximated by the square root of the total count). When cells are touching a line, it is common practice to count cells that touch the top and left lines of each square, and not to count cells that touch the bottom and right lines. The hemocytometer can also be used to distinguish living cells from dead cells and this can be used to calculate the percentage of viable (i.e., living) cells in the culture. For this purpose a "vital stain" is used. A vital stain is a dye that living cells are able to exclude from their interiors. Dead and dying cells can not exclude the dye and therefore show signs of color. For this purpose the most commonly vital stain is trypan blue. The percentage of viable cells in a culture is calculated as number of living cells / (number of dead cells + number of living cells) × 100. It is therefore important to keep two tallies while counting cells in a hemocytometer, the number of cells that have dye and the number that do not. Procedure Counting Non-Adherent Cells T27A is a very aggressive murine (mouse) leukemia cell line that is non-adherent and grows as a suspension culture. As you will see, the growth characteristics of normal cells and cancerous cells are very different. Normally these procedures would be carried out under a laminar low hood (see Chapter 12) to avoid contamination by bacteria and fungi. However, since today's exercise only involves counting cells, and not subculturing or reincubating them, it is possible to work on the open bench under nonsterile conditions. 1. Obtain a culture of T27A cells (in the incubator in a blue-capped culture flask). 2. Remove 1 ml of cell culture and place it in a 2 ml microcentrifuge tube. 3. Centrifuge the cell suspension for 5 min at 2000 rpm in the bench top centrifuge. 4. With a Pasteur or transfer pipette, remove the medium from over the cell pellet. Be careful not to disturb the cell pellet. 5. Add 100 l of phosphate buffered saline (PBS) to the cell pellet and resuspend the cells by drawing them into the pipette tip several times. 6. To a small test tube add: 25 l of the PBS/cell suspension just prepared 25 l 0.04% trypan blue solution wait 30 seconds then add 50 l PBS

7. Mix the suspension by vortexing, swirling, or flicking with your finger several times.

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8. Add a small amount of the cell mixture to the hemocytometer. Use low power to locate the grid in the microscope. Switch to a higher power, one that just allows the center grid (the one labeled "A" in Figure 2) to fit into the field of view From this point there are two ways to count cells and calculate cell densities using a hemocytometer. The method you choose depends on the number of cells present. 8a. If the number of cells visible in the large central square (square "A") is less than about 200 (make a ball-park estimate), count the number of cells in the entire large square. Then count the number of cells in the four corner squares (the squares touching the corners of square "A"). Count the number of both living (cells that exclude dye) and dead cells (cells that appear blue). For the most accurate count, count more than 200 cells, but do not stop counting until the count of a given square is complete. Switch to the other side of the hemocytometer and repeat the counts. Recall that with the cover slip in place, the volume of one large square is 1 ×10-4 mL. 9a. The cell density is calculated as: The # of cells counted / number of squares counted × 104 × dilution factor = number of cells / mL in the original culture. Cell viability is calculated as: Number of living cells ÷ (number of dead cells + number of living cells) × 100 = percent viability. The dilution factor (or concentration factor, as it could be called if you concentrate the cells by centrifugation) is the factor you must multiply your cell density counts by in order to correct for any dilutions (or concentration) you made. The dilution factor can be calculated as (the volume you end up with) / (the volume you started with). For example, if you mix together 1 mL of cell suspension and 1 mL of trypan blue, the cell density you obtain will be one half that in the original culture because the cells have been diluted in half. In this case the dilution factor is 2/1 = 2, your counts must be multiplied by two. On the other hand if you take 1 mL of cell suspension, pellet the cells by centrifugation, then bring them up in 100 L, you have concentrated the cells 10-fold. The dilution factor in this case (now a concentration factor) is 0.1/1.0 = 0.1. Your counts must be multiplied by 0.1 (i.e., divided by 10) to correct for the concentration. 8b. If a very large number of cells are visible, count the cells in each of the twenty-five small squares located within the larger square "A". Count the number of both living (cells that exclude dye) and dead cells (cells that appear blue). For the most accurate count, count more than 200 cells, but do not stop counting until the count of a given (small) square is complete. Otherwise count all 25 small squares in the central grid ("A"). Using the stage adjuster, move to the other counting chamber and count the number of cells in the central grid of that side. 9b. Calculate the cell density and cell viability of the original culture. a. Cell density: Number of cells counted ÷ number of small squares counted (in the central grid) ÷ 10 = 106 cells / ml of original culture. b. Cell viability. Number of living cells ÷ (number of dead cells + number of living cells) × 100 = percent viability.

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10. Repeat the process on the same flask of cells to determine your reproducibility. Counting Adherent Cells Many cell types, including the mouse melanoma line B16-F1 and the CHSE and ZF-L fish cell lines, grow in culture by adhering to the substratum (i.e., the culture flask). In order to count these types of cells they must be enzymatically detached before they can be counted. Follow the procedure below to count these cells. Once again, because we are not going to continue to culture these cells (all the cells we use will be discarded), counting can be performed under nonsterile conditions. 1. Remove the cell culture medium with a pipette. 2. Add 5 mL of calcium- and magnesium-free PBS. Wash the cells by gently swirling the PBS around the bottom of the flask. 3. Remove the PBS, then add 2 mL of trysin/versene solution. 4. Allow the cell culture to digest for 5 minutes or more, until most of the cells have become detached from the bottom of the flask. Watch the cells carefully under the inverted microscope! Do not allow the digestion to proceed for too long, cell might be damaged. 5. Add 5 mL of fresh culture medium and mix the suspended cells thoroughly but gently. 6. Remove 1 ml of cell culture and place in a 2 ml microcentrifuge tube. 7. Centrifuge the cell suspension for 5 min at 2000 rpm in the bench top centrifuge. 8. With a Pasteur pipette, remove the medium from over the cell pellet. Be careful not to disturb the cell pellet. 9. Add 100 l of PBS to the cell pellet and resuspend the cells by drawing them into the pipette tip several times. 10. To a small test tube add: 25 l of the PBS/cell suspension just prepared 25 l 0.04% trypan blue in PBS wait 30 seconds then add 50 l PBS 11. Mix the suspension by vortexing or swirling. 12. Add a small amount of the cell mixture to the hemocytometer and decide which counting method to use based on the criteria present above (Steps 8 and 9).

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13. Count the number of both live cells (cells that exclude the dye) and dead cells (appear blue). For the most accurate count, count more than 200 cells. Using the stage adjuster, move to the other counting chamber and count the number of cells on that side. 13. Calculate the cell density and cell viability of the original culture.

Class Comparisons Repeat the procedures above on at least three different cultures of both adherent and nonadherent cells. Write your results on the board in the appropriate space and compare your results to those of others who have counted the same flask. Questions. 1. How do you counts compare to those of others? 2. Using what you know about the physical dimensions of the counting chambers and the dilutions you made to the cell cultures, confirm the calculations used in steps 9a and 13a above. References Harrison, R.G. 1907. Observations on the living developing nerve fiber. Proc. Soc. Exp. Biol. Med. 4:140-143.

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Chapter 11.

Cell Culture: Primary Culture of Chick Embryo Fibroblasts.

___________________________________

Objective To establish primary cultures of fibroblasts from chicken embryos. Introduction In this week's exercise, you are going to use a chicken embryo to establish a primary cell culture of fibroblasts. Of course, the chicken embryo contains many types of cells, and your initial culture will therefore be a heterogeneous collection of cell types. Within a few hours, however, the culture will become dominated by fibroblasts. These cells are particularly hardy and tend to overgrow other adherent cells types. Fibroblasts are connective tissue cells that are flat and elongated with many cytoplasmic processes extending from the cell body. They are particularly abundant just beneath the skin of adult vertebrates, and in embryos they tend to be found throughout the body. Fibroblasts are the cells that form the support structure for many tissues and organs. They normally differentiate into cell types that include chondroblasts, collagenoblasts, and osteoblasts, and form the fibrous tissues in the body including bones and tendons. Primary cultures are defined as those derived directly from excised, normal animal tissue. Primary culture are grown either as an explant culture, that is as a relatively intact piece of tissue, or as single cells after dissociation into a cell suspension by enzymatic digestion. In most cases, the preparation of primary cultures is more labor intensive than growing secondary cultures, mainly because of the need for obtaining and caring for the original animal and the time and material necessary to prepare the tissue. Normal cells, that is cells that are not stem cells or cancerous cells, can be maintained in vitro only for a limited period of time. During their relatively limited life span, primary cells usually retain many of the differentiated characteristics that the cell would have in the originating organism. Cancer cells and some stem cells can be propagated indefinitely because they do not undergo replicative senescence. These are called immortal cells for that reason. Stem cells grow and divide indefinitely because their normal function is to replace damaged or aged cells in their tissue of origin. For example, adult stem cells routinely replace the lining our digestive system and continuously produce red blood cells. Cancer cells grow indefinitely because they have lost the ability to limit their own growth, as normal cells do. Cancer cells are often derived from actual clinical tumors, but transformation may also be induced using viral oncogenes or by chemical treatments. Transformed cell lines present the advantage of almost limitless availability, but the disadvantage of having retained very little of the original in vivo cell characteristics. Sterile Techniques Sterile (or aseptic) techniques are used to prevent contamination, specifically biological contamination, of your cultures. Life forms other than those you intend to grow will readily grow in your culture medium unless you prevent them form doing so. In this laboratory, and in all laboratories where cells of metazoans are grow, unwelcomed cells include bacteria, fungi and

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viruses. These uninvited visitors exist everywhere in the air and on the surfaces of all objects. They can fall into cultures and can be introduced via contaminated utensils and media. A conscious effort must be made to keep them out of a sterile environment. Sterile techniques therefore focus on covering and sterilizing all utensils (e.g., pipettes), media, culture vessels and the working area, and then keeping contaminating microorganisms away. Even though antibiotics are included in many media preparations, they can not be counted on to mitigate gross contamination resulting from poor sterile technique. A major problem in culturing cells in this class is contamination caused by carelessness and improper techniques. Proper sterile techniques are common sense and become second nature once the arguments above are understood. Some common practices you should always follow include: 1. When possible, work in the laminar flow hood. These hoods (described in Chapter 13) are designed to prevent airborne contamination of cultures and media. 2. Always swab work surfaces with 70% ethanol before beginning and when finished for the day. Also, swab during your work if you feel it is necessary. 3. Swab any instruments that will be used in the hood with 70% ethanol, particularly the pipettor handles and pump tubes, which will often be used above biological samples. 4. Never open a sterile container unless it is under the hood. 5. Minimize the time that containers are left open. Close or cover them when not in use. 6. Loosen the lids and caps of all containers but do not remove them until you are ready to use that container. 7. After opening a container, do not place the cap on the working surface. Instead, hold on to it (between your fingers if necessary) and keep it facing down. If you must put it down, swab an area in the rear of the hood (where there is less traffic) and place the cap in this area, facing down. 8. Use a separate sterile pipette for each manipulation. Never draw from a flask or bottle with a pipette that has already been used elsewhere for another manipulation. (Except in cases where you are certain that no contamination will result. For example, under some circumstances it may be acceptable to draw from a bottle of fresh medium, deliver this to a flask of cells and then to use the same pipette to distribute those cells into new flasks). When in doubt, use a new pipette. 9. Do not exhale directly into your culture while working under the hood. 10. Never pour from one sterile container into another. Always use a pipette. Pouring can leave a path of liquid from the interior of the container to the exterior that can introduce contamination.

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11. Clean up drips and spills immediately and swab the area with 70% ethanol. 12. Avoid contact between the tip of the pipette and the mouth of the bottle. The mouth and neck of the bottle (both inside and out) present a potential sources of contamination. 13. Leave sterile pipettes in their wrapping until just before use. 14. The best rule of thumb is to always be thinking of sterility. Much of our work will be done at the bench and not in the hood, so be mindful of contamination. 15. When finished, always meticulously clean up the work surface for the next user. Put trash in the appropriate receptacle, glassware in the wash area by the sink, and swab the area with 70% ethanol. You expect to find the area clean when you arrive, so leave it that way for the next person. Cell culture media and CO2 incubators Culturing cells requires not only specific techniques, but also some specialized equipment and materials. Among the most important are the culture medium and CO2 incubators for maintaining the cultures. Media. Media for cell culture can be prepared in the lab, but it is far more efficient to purchase media from commercial suppliers. These media are inexpensive and are of very high quality. They often have odd names that memorialize the original formulator (e.g. Ham's F-10, Eagle's Medium, Dulbecco's Modified Eagle's Medium, Liebovitz Medium) or place of origin (RPMI 1640, after "Roswell Park Medical Institute"). Other examples include L-15, Medium 199, and IMR-90. Though formulated for different and specific cell culture needs, all of these media have in common some ingredients. All are composed of aqueous solutions of inorganic salts and buffering agents and contain various proportions of carbohydrates, amino acids, vitamins, fatty acids, other lipids, proteins and peptides and serum. The inorganic salts help to retain the osmotic balance of the cells and help to maintain normal cell electrical potentials by providing sodium, potassium, calcium, magnesium, and other ions. Some of these are required for cell attachment to the substratum, others function as cofactors for enzymes. Buffering systems are required because most cells function properly only within well defined limits of pH, usually between 7.0 and 8.0. Fibroblasts prefer pH of 7.4 to 7.6 for optimal growth whereas some immortal cell lines prefer pH in the range of 7.0-7.4. The regulation of the pH of the culture medium is particularly important immediately following cell seeding when a new culture is establishing. Cell culture media are buffered in one of two ways. One is by what is called a "natural" buffering system in which CO2 provided by the incubator (see below) balances with the normal CO2 / bicarbonate content of the culture medium. The other method uses chemical buffering agents such as HEPES, MOPS, or any of numerous others. Cultures using natural bicarbonate/CO2 buffering systems need to be maintained in an atmosphere of 5-10% CO2 in air. Chemically buffered media are also held under the same conditions in order to mimic the normal environment of the cell. Bicarbonate/CO2 is low cost and non-toxic. Chemical buffering agents have superior buffering capabilities but are more expensive and can be toxic to some cell types at higher concentrations. Most commercial culture media also include phenol red as a pH indicator so that the pH status of the medium can be instantly monitored simply by observing the color. Phenol red turns yellow when acidic an purple when alkaline. Usually the

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culture medium should be changed if these colors are observed. Carbohydrates are the major source of energy for cells in culture. The most common one used in cell culture media is glucose, but galactose, fructose, and maltose are also used. Vitamins are required to serve as cofactors for enzymes. The B vitamins, especially B12, are necessary for the growth of some cell lines. Proteins, peptides, and lipids help simulate the normal cell environment. The most common proteins used are albumin, transferrin, and fibronectin, and free fatty acids and cholesterol are also sometimes included in cell culture media. Perhaps the most important component of cell culture formulations is serum. Serum is a very complex mixture of albumins, growth factors and other essential ingredients that is virtually impossible to mimic at the lab bench. The most commonly used serum is fetal bovine serum, and calf and horse sera are also used. An advantage of these sera is that they are readily available and can be used on a wide variety of cell types. Growth factor receptors are highly conserved between species. For example, murine epidermal growth factor can stimulate the growth of fish cells in culture. Depending on the type of culture, other sera may be used. For instance, fish cell cultures grow best in the presence of fish serum. CO2 incubators. Most mammalian cells are grown in an atmosphere of 5-10% CO2 in air because this is the CO2 concentration of the blood and tissues of those organisms. Fish cells require CO2 in the range of 1-2 % for the same reason. Obviously, CO2 is an absolute requirement in cell cultures using the bicarbonate/CO2 buffering system. Culture flasks should have loosened caps to allow for sufficient gas exchange. Cells should be left out of the incubator for as little time as possible and the incubator doors should not be opened for very long. The humidity must also be maintained for cultures growing in tissue culture flasks with loose caps, so a pan of water is kept filled at all times. When using CO2 incubators, always be aware of the CO2 level in the bottle supplying the gas (shown on the gauge on the first stage of the regulator), and the water level of the humidifier pan. Procedure A. Removal of the embryo. 1. Swab the large end of an egg with 70% ethanol. 2. Carefully puncture the large end of the shell with the point of a pair of sterile scissors. Using plastic forceps, break the away a circle of shell around the air sac to expose the underlying membrane (the chorioallantois). 3. Remove the chorioallantoic membrane, exposing the embryo. 4. Pour the embryo into the deeper half (the bottom) of a sterile Petri dish. 5. Pour 12 mL of sterile HBBS into the other half of the Petri dish. 6. Remove the head of the embryo with forceps. Transfer the body to the HBSS and rinse it to remove yolk and blood cells. 7. Transfer the washed carcass to a sterile Erlenmeyer flask with a stir bar.

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B. Preparation of cell suspension. 8. Add 5 mL of trypsin/versene solution to the flask and stir the mixture on a low setting on a magnetic stir plate. Allow to digest for three minutes. 9. Meanwhile, add 20 mL of CEF culture medium to a 50 mL centrifuge tube. 10. After three minutes of digestion, remove the flask from the stir plate, allow the tissue to settle, then tilt it slightly and pipette off the cell suspension (free of any large tissue fragments). Transfer the cell suspension to the medium in the centrifuge tube. 11. Add an additional 5 mL of trypsin/versene to the pieces of tissue remaining in the flask and again stir for three minutes. After digestion, remove the cells suspension as you did previously (in step 10) and add it to the culture medium in the centrifuge tube. 12. Repeat step 11. 13. At this point in the procedure you should have a centrifuge tube containing 20 + 5 + 5 + 5 = 35 mLs of cell suspension. 14. Centrifuge the cell suspension at 500 × g for 5 minutes. 15. Decant the supernatant and resuspend the cell pellet in 5 mL of fresh CEF culture medium. C. Determination of the cell density of the suspension and starting the primary cultures. 16. Using a sterile pipette, transfer 0.1 mL of cell suspension to a small glass test tube. 17. Add 0.5 mL of trypan blue solution and 1.4 mL of HBSS (What is the dilution factor now? Dilution Factor = _________. ) 18. Following the procedures you used last week for loading the hemocytometer and counting cells (Chapter 10), but using HBSS as diluent (instead of PBS), determine the cell density of your suspension. Remember to count both viable and non-viable cells, and to count both chambers to get duplicate counts. 19. Calculate the volume of the cell suspension that contains 6 × 105 cells. With a sterile tip, seed that volume into two sterile 25 cm2 culture flasks. 20. Add 5 mL of CEF culture medium to the flasks (use a sterile pipette). Leave the caps loose (or set the "Y" on the cap to the ventilate position) so gases can exchange. 21. Label your flasks and place them in the CO2 incubator. 22. Check your flasks every day using the inverted microscope and note cell shape and estimate confluence (the percent of the bottom of the flask that is covered with cells).

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References Chipev, C. C. and M. Simon. 2002. Phenotypic differences between dermal fibroblasts from different body sites determine their responses to tension and TGF1. BMC Dermatology 2:13-20. Freshney, R. I. 1983. Culture of Animal Cells: A Manual of Basic Technique. Alan R. Liss, Inc. New York. Mischell, B. B. and S. M. Shiigi. 1980. Selected Methods in Cellular Immunity. W.H.Freeman & Co. San Francisco. Rothblat, G. H. and V.J. Cristofalo, eds. 1972. Growth, Nutrition and Metabolism of Cells in Culture. Academic Press, New York. Spierenberg, G.T., et al. 1984. Phototoxicity of N-2-hydroxyethylpiperazine-N-ethanesulfonic acid-buffered culture media for human leukemic cell lines. Cancer Research 44: 22532255. Wang, R.J. 1976. Effect of room fluorescent light on the deterioration of tissue culture medium. In Vitro 12:19-22. Zigler, J.S., et al. (1985) Analysis of the cytotoxic effects of light-exposed HEPES-containing culture medium. In Vitro 21: 282-285.

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Chapter 12.

Cell Culture: Secondary Cultures of Chick Embryo Fibroblasts.

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Objective To passage primary cultures of chick embryo fibroblasts in order to establish secondary cultures, and to then monitor the confluency, growth rate, and viability of the secondary cultures over one week. Introduction Last week you prepared primary cultures of fibroblasts from chicken embryos. This week's exercise involves subculturing the primary cultures into secondary cultures and carefully monitoring the confluency, growth rate, and viability of the secondary cultures. As you recall, cell cultures established from animal tissues are called primary cultures. The cells from such cultures can be removed from the flask and used to establish a large number of secondary cultures. Cells from the secondary cultures can then be used to establish more and more cultures until the cells undergo replicative senescence. The terms used to describe this repeated transfer of cells are passaging, subculturing, seeding, and cell-plating. All these terms are synonymous. "Eyeballing" Cell Cultures Before continuing with any procedure That uses cell cultures, the overall heath of the cultures should first be visually inspected using a practice celled "eyeballing". This can be done quickly and quantitatively by making the following observations: 1. First, the culture medium should be transparent. A cloudy medium often indicates contamination by bacteria or fungi. 2. Check the pH of the culture medium by observing the color of the pH indicator (phenol red). As a culture becomes more acid, the indicator shifts from red to yellow-red to yellow. As the culture becomes more alkaline, the color shifts from red to purple (actually fuchsia, or purpleish-redish. Most cells tolerate slightly acid conditions better than they tolerate alkaline. As you have already learned, observing living cells in culture flasks requires a special microscope. The phase contrast optics of these microscopes enhance the contrast between the background and living cells, making them much easier to see. Use the inverted scope to check the following: 3. If the culture contains adherent cells, check for cell attachment. The vast majority of cells should be attached to the substratum and spread out in a shape characteristic of the cell type. 4. Check the shape of the cells. Are the cells their characteristic shape? Or do they have a rounded or irregular appearance? If cell shape is not "normal", the cells are probably not doing well. 5. Look for giant cells. The number of giant cells will increase as a culture ages or becomes less healthy. These should be very rare in newly established, healthy cultures. 6. Check for "floaters". Floaters are cells that are not attached to the substratum, and

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thus are floating in the medium. There are two reasons for adherent cells to be floating, they are dividing or they are unhealthy. A healthy culture will normally have a small percentage of floaters because cells must detach from the substratum before undergoing mitosis. In this case, floaters are a sign of a healthy culture. On the other hand, large numbers of floaters probably indicate a culture in poor health. 7. Check for confluence. The growth of a cell culture can be estimated with some precision by assessing the percentage of the culture flask covered by cells on consecutive days. By comparing the amount of space covered by cells with the unoccupied spaces, the percent confluency can be readily estimated. 8. Check for the presence of vacuoles. An unusually large number of vacuoles, visible as clear spherical "bubbles" within the cells, is a sign of an unhealthy culture. Try replacing the culture medium and checking again after 24 hours. 9. A good indicator of the health of a cell culture is the rapidity at which the cells attach and spread out once they have been passaged. Attachment within an hour or two suggests that the cells are healthy, have not been traumatized, and that their environment is not grossly abnormal. Longer attachment times are suggestive of problems. However, good cultures can result even if attachment does not occur for several hours. 10. Tumor cells do not stop growing when confluent and therefore tend to pile-up on one another. If this is observed, the culture should be passaged.

Procedure A. Subculturing primary cultures. 1. Eyeball the cultures and note any irregularities. 2. With a sterile pipette, remove the culture medium from the flask. (Dispense it into a waste Erlenmeyer flask.) 3. Add 3 mL of Ca2+ and Mg2+ -free phosphate buffered saline, rock the flask several times to completely remove any remaining culture medium, then remove the saline. 4. Add 1 mL of trypsin/versene to the monolayer. Ensure the entire surface is covered with solution. 5. Using the inverted microscope, carefully monitor the cells every few minutes. The cells will become more and more spherical as they detach from the substratum. 6. When most (90-99%) of the cells have detached, add 5 mL CEF culture medium. Do not allow the trypsin to work longer than it takes to detach approximately 90% of the cells. 7. Aspirate the cell suspension several times in and out of a sterile pipette in order to break apart cell aggregates and to mix the suspension. 8. Add 0.1 mL of cell suspension and 0.1 mL of trypan blue solution to a small glass test tube. Record the dilution factor (Dilution Factor = _________).

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9. Following the procedures you have used previously (Chapter 10), determine the cell density of your suspension. Remember to count both viable and non-viable cells, and to count both chambers to get duplicate counts. 10. Calculate the volume of the cell suspension that contains 2 × 105 cells. With a sterile tip, seed that volume into 4 sterile 25 cm2 culture flasks. 11. Add 5 mL of CEF culture medium to the flasks (use a sterile pipette). Leave the caps loose (or set the "Y" on the cap to the ventilate position) so gases can exchange. 12. Label your flasks and place them in the CO2 incubator. B. Determination of Confluency, Growth Rates, and Viability of Secondary Cultures On days 1, 3, 5, and 7 (day 7 is the next lab period), carry out the following procedures: 1. Eyeball the cultures and note any irregularities. 2. Using the inverted microscope, estimate cell confluency (percent of substratum covered by cells). 3. Follow steps 2-9 in Part A above and determine the cell density and cell viability of each of your cultures on their respective day. C. Data Reduction and Analysis. 1. Using the data you have collected over the past week, plot the following: a. The estimated confluence as a function of culture time. b. The cell density as a function of culture time. c. Cell viability as a function of culture time.

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Chapter 13.

Cell Culture: Tumor Cell Cultures.

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Objective To passage and propagate a tumor cell line, and to then monitor the growth rate and viability of the culture over one week. Introduction Despite recent advances in the diagnosis and treatment of cancer, tumor cell progression and metastasis constitute a major cause of death in cancer patients. Cancer results when the growth of cells becomes uncontrolled and uncontrollable. By definition, cancer cells are immortal. Because their growth is unregulated, cancer cells propagate rapidly as long as the appropriate nutrients are available. The unrestrained expansion of cells is what eventually kills the organism the cells inhabit. A cancer cell is formed when a normal cell acquires a mutation that causes a growth regulatory pathway to fail. A variety of mechanisms can cause this "transformation" (cancer cells are often called transformed cells). Ultraviolet and X ray radiation, viruses, and chemical carcinogens can all cause mutations in DNA. Some chemical compounds are not carcinogenic themselves, but are converted into carcinogens by the organism's metabolism. Spontaneous mutations can also. If the mutation is not corrected by one of the myriad DNA correction enzymes, it could lead to altered genetic activity and cancer. Regardless of how a normal cell is transformed, some type of stable gene alteration must occur. The mutation causes the abnormal gene to produce an abnormal protein product. Two categories of genes play major roles in initiating cancer. In normal cells, these genes and their associated proteins control some aspect of progression through the cell cycle. One category of genes, the proto-oncogenes, are encourages cell division. The other category, called tumorsuppressor genes, inhibits it. Together, proto-oncogenes and tumor-suppressor genes coordinate the regulated growth that normally ensures that tissues and organs maintain an appropriate size and morphology. Mutations in either can cause cancer A proto-oncogene is a gene in normal cells that functions during normal cell growth. Most proto-oncogenes code for proteins that are involved in the molecular pathways that receive and process growth-stimulating signals from other cells in a tissue. Typically, such signaling begins with the production of a growth factor. A stable mutation can convert the proto-oncogene into a oncogene. Expression of the oncogene results in cell transformation. In general, oncogenes cause the proteins involved in growth-promoting pathways to be overactive, and the cell proliferates much faster than it would if the mutation had not occurred. Some oncogenes cause cells to overproduce growth factors. Others produce aberrant receptor proteins that convey stimulatory signals into the cytoplasm even when no growth factors are present in the environment. Still others disrupt signaling cascades such that the nucleus receives stimulatory messages continuously, even when growth factor receptors are inactive. There are also oncogenes that code for defective protein kinases and transcription factors. Mutations in tumor suppressor genes appear to cause more than half of all human

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cancers. In normal cells, growth stimulating signals are balanced by growth inhibiting signals, and many of these come from tumor suppressor genes. Mutations in tumor suppressor genes can cause cells to become cancerous by allowing them to become resistant to the inhibitory messages. The normal functions of many known tumor-suppressor genes are still unclear. Sterile Cell Culture Hoods There are two kinds of sterile hoods, vertical and horizontal. Both types can be found in the Department of Biological Sciences, but in this class we will use a vertical hood for cell culturing. In both types of hood, air is filter by passage through a HEPA (high efficiency particle) filter that removes particulates. In vertical hoods, also called biology safety cabinets, air is forced down from the top of the cabinet through vents located in the surface and back of the work area before being filtered and released into the environment. In horizontal hoods, air is filtered then blown from the back of the cabinet out at the operator in a horizontal fashion. Vertical hoods are preferred, and sometimes required, for working with hazardous organisms because the aerosols that are generated in the hood are filtered out. Horizontal hoods are not suitable for work with hazardous organisms because the air is forced directly onto the user, but these hoods offer superior protection against contamination. It is extremely important to understand that sterile hoods are not fume hoods (like that found in the back of the Cell Lab). Thought they look similar, fume hoods and sterile hoods have profoundly different functions. Sterile hoods are for use with hazardous, volatile, and explosive chemicals. They should never be used for cell culture work. Conversely never use chemicals, other than those used in normal cell culture techniques, in the sterile hood. Essential practices when using sterile hoods include the following. 1. Swab down the work surface liberally with 70% ethanol. Start from the back and proceed forward. 2. Swab while working, and immediately after wiping up drips and spills. 3. Boxes of sterile pipette tips should remain in the hood and not be taken out. Once taken out of the hood and opened, they are no longer sterile. The ultimate transgression is to take the box out of the hood, open it, then return it to the hood, so that the next users think they are sterile. Never do this. Leave the tips in the hood. 4. Containers should always be tightly capped when outside the hood. That means they should have been tightly capped the last time they were in the hood. 5. Thoroughly dry containers that have been warming in the water bath, then swab them with 70% ethanol (particularly the cap, neck, and bottom) before placing them in the hood. 6. Place the items you need to carry out a particular procedure (bottles of media, pipettes, tips, waste beaker, etc...) into the hood before you begin work. Put in the hood only the things you need. This will reduce the number of times you will need to leave and then reenter the hood and minimize cluttering in your work space. Keep nonessential and infrequently used items at the sides of the hood and your immediate working materials in a clear working space in the center of the hood. 7. Avoid blocking the air vents at the front and back of the hood. 8. Work near the center of the hood. Avoid working too close to the fron whre contamination is more likely. 9. Confirm that items that should be autoclaved actually have been by noticing that the stripes on autoclave tape turn black after heating. If an item has autoclave tape on it, do not use it unless the color change has occurred.

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In this exercise, you will culture and propagate mouse tumor cells and assess their growth rate. The growth rate of the tumor cells will be compared to that of the chick embryo fibroblasts. The cell line you will be using is the B16-F1 strain of murine melanoma. This is an adherent cell line that grows, morphologically, very much like chicken fibroblasts.

Procedure Pre-warm all media (except trypsin) before use and perform all work in the large hood in the cell culture room. 1. Obtain a flask of B16 cells and remove the cell culture medium with a sterile pipette. 2. Add 3 mL of calcium- and magnesium-free PBS. Wash the cells by gently swirling the PBS over the cell monolayer. 3. Remove the PBS (again with a sterile pipette), then add 1 mL of trysin/versene solution. 4. Allow the cell culture to digest for 5 minutes or more until most of the cells have become detached from the bottom of the flask. Watch the cells carefully under the inverted microscope while treating with trypsin! Do not allow the digestion to proceed for too long or cells may be damaged. 5. Add 5 mL of fresh B16 culture medium and mix the suspended cells thoroughly but gently by aspirating them up and down a few times in the pipette. 6. Remove 0.1 ml of cell culture and place in a small glass test tube. 7. Add 0.1 mL of trypan blue solution and mix by flicking the tube with a finger tip. 8. Add a small amount of the cell mixture to the hemocytometer and decide which counting method to use based on the criteria presented previously (see Chapter 10). 9. Count the number of both living and dead cells. Calculate the cell density and cell viability of the original culture. 10. Calculate the volume of the cell suspension that contains 2 × 105 cells. With a sterile tip, seed that volume into 4 sterile 25 cm2 culture flasks. 11. Add 5 mL of B16 culture medium to the flasks (use a sterile pipette). Leave the caps loose (or set the "Y" on the cap to the ventilate position) so gases can exchange. 12. Label your flasks and place them in the CO2 incubator. B. Determination of Growth Rates and Viability of Tumor Cell Cultures On days 1, 3, 5, and 7 (day 7 is the next lab period), carry out the following procedures:

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1. Eyeball the cultures using the inverted microscope and note any irregularities. 2. Follow steps 3-9 in Part A above and determine the cell density and cell viability of each of your cultures on their respective day. C. Data Reduction and Analysis. 1. Using the data you have collected over the past week, plot the following: a. The cell density as a function of culture time. b. Cell viability as a function of culture time. D. The Lab Report. In your laboratory report, compare and correlate graphically the growth rate of the CEF cells and the B16 as determined by the cell counts and your simultaneous estimate of confluence. In your discussion, draw conclusions about your observations and relate them to the life histories of these cells.

References These references describe the B16 cell line. Algarra, I., M.Perez, P.Hoglund, J.J.Gaforio, H.G.Ljunggren, and F.Garrido. 1993. Generation and control of metastasis in experimental tumor systems; inhibition of experimental metastases by a tilorone analogue. International. Journal of Cancer 54:518-523. Calorini, L., A.Fallani, D.Tombaccini, G.Mugnai, and S.Ruggieri. 1987. Lipid composition of cultured B16 melanoma cell variants with different lung-colonizing potential. Lipids 22:651-656. Korycka, B.M. and R.P.Hill. 1989. Dynamic heterogeneity: experimental metastasis studies with RIF-1 fibrosarcoma. Clinical & Experimental. Metastasis 7:107-116. Kramer, R.H. and G.L.Nicolson. 1979. Interactions of tumor cells with vascular endothelial cell monolayers: a model for metastatic invasion. Proc. Natl. Acad. Sci. U. S. A. 76:5704-5708. Trulla, L.L., A.Magistrelli, M.Salmona, and M.T.Tacconi. 1993. Effect of cell density on cytotoxicity of ether lipid analogues in variants of B16 murine melanoma. Lipids 28:403406.

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Appendix 1. Reagents and Materials.

Chapter 2. Pipetting and Creation of a Standard Curve. Apparatus Spectrophotometers (8) Micropipettors and tips Materials a) Test tubes, 12 x 75 mm b) Test tube racks c) Protein assay reagent (from BioRad) d) Protein stock solution (BSA, 2mg/mL) e) "Unknown" protein solutions, diluted so that 100 µL contains protein in the rage of the standard curve. Chapter 3. Cell Fractionation: Isolation of Mitochondria from Cauliflower. Media, Supplies, Apparatus. Cauliflower (one head per section) Triple beam balances (2) Isolation buffer (0.3 M D-mannitol, 0.02 M phosphate buffer, pH 7.2). Make 2.0 L for 4 sections. Assay buffer (0.3 M D- mannitol, 0.01 M KCl, 0.005 M MgCl2, 0.02 M phosphate buffer, pH 7.2). Make 2.0 L for 4 sections (for this week and next). Oak Ridge tubes (35 mL) Sea sand Cheesecloth 50 mL centrifuge tubes 15 mL centrifuge tubes razor blades 70% ethanol (750 mL 95% EtOH up to 1000 mL with dH2O). Mortar and pestle, stored in the freezer overnight Pre-chill the centrifuges Large weigh boats Beakers for ice Ice Isolation buffer. D-mannitol 27.3 g ×4= 109.2 g KH2PO4 0.4 g ×4= 0.16 g 1.2 g ×4= 4.8 g K2HPO4 Distilled H2O 400 mL ×4= 1600 mL Adjust pH to 7.2 with 1.0 M KOH, bring the volume up to 500 mL with distilled water using a volumetric flask. Store at 4°C. Multiply all weights and volumes by 4 to make 2 L.

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Assay Buffer. D-mannitol 27.3 g ×4= 109.2 g 0.4 g ×4= 0.16 g KH2PO4 K2HPO4 1.2 g ×4= 4.8 g KCl 0.4 g ×4= 0.16 g MgCl2-6H2O 0.5 g ×4= 0.20 g Distilled H2O 400 mL ×4= 1600 mL Adjust pH to 7.2 with 1.0 M KOH, bring the volume up to 500 mL with distilled water using a volumetric flask. Store at 4°C. Multiply all weights and volumes by 4 to make 2 L.

Chapter 4. Cell Fractionation: Assay of Mitochondrial Enzyme Activity. Spectrophotometers 13 x 100 mm test tubes 1 mL and 5 mL pipettes (bulk pack) Pasteur pipettes Assay Buffer (see above) 0.04 M sodium azide 50 µM DCIP 0.2 M succinate Thaw the postmitochondrial and mitochondrial pellets collected last week 0.4 M sodium azide Bring 0.26 g of sodium azide up to 100 mL distilled water in a volumetric flask. Store at room temperature. 50 µM DCIP Place 7.25 mg aliquots of 2,6-dichlorophenolindophenol (Na salt) into clean 50 mL centrifuge tubes (one for each section). Just before use, add 50 mL distilled H2O to a tube and vortex thoroughly. 0.2 M succinate Bring 5.4 g of sodium succinate-6H2O up to approximately 90 mL, adjust the pH to 7.0 with 1 M HCl, bring up to a final volume of 100 mL using a volumetric flask. Store at 4°C.

Chapter 5. Cell Fractionation: Enzyme Specific Activity. Requires a tube of "mitochondria" from Chapter 4. Store in freezer until this week. Apparatus Spectrophotometers Micropipettors and tips

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Supplies BioRad protein assay reagent, dilute (1:5) 50 mL / student pair Protein standard, 2mg/mL transfer pipets, plastic 12 mm × 75 mm culture tubes PBS

Chapter 6. Enzyme Kinetics. Apparatus Spectrophotometers Micropipettors and tips Supplies transfer pipets, plastic Pasteur pipets and bulbs 12 mm × 75 mm culture tubes 50 mL centrifuge tubes 15 mL centrifuge tubes 10 mL pipettes 1 mL pipets Reagents Alchol dehydrogenase (ADH). Stock is 60 units / mL , make 6 mL. Working solution: Dilute 1.2 mL stock with 22.8 mL dH2O. 24 mL is good for 1 section. 3M ethanol For each 100 mL desired (200 mL is good for four sections), mix 18.5 mL 95% ethanol 81.5 mL dH2O Pyrophosphate buffer (0.15 M pyrophosphate buffer with 0.036 M -mercaptoethanol, pH 8.5) Stock 0.15 M sodium pyrophosphate Add 16.7 g Na4P3O7 ­ 10 H2O (MW = 446.06), to dH2O and bring up to 250 mL. Stock 0.15 M phosphoric acid Add 2.55 mL of phosphoric acid (85% H3PO4) to 200 mL dH2O. Bring up to a total volume of 250 mL. To make 200 mL of final pyrophosphate buffer:

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Add stock phosphoric acid solution drop-wise to 50 mL stock sodium pyrophosphate solution until the pH reaches 8.5. TO this add 0.5 mL of mercaptoethanol and bring the mix to a final volume of 200 mL. 0.015 M -NAD+ (MW = 663.5) For each 25 mL desired, bring 249 mg -NAD+ to a total volume of 25 mL. Add one frop opf 0.1 N NaOH to stabilize the solution. (this will remain stable for several days at 4°C)

Chapter 7. Protein Fractionation: Isolation of IgG from Human Serum. Apparatus Spectrophotometers Micropipettors and tips Supplies Dialysis tubing Human serum DEAE cellulose Bovine serum albumin BioRad protein assay reagent 50 mL centrifuge tubes, 2 / pair 15 mL centrifuge tubes, 3 / pair transfer pipets, plastic Pasteur pipets and bulbs 12 mm × 75 mm culture tubes From the new Manual: Part A. Dialysis of human serum. 1. Dialyze 30 mL of human serum against 0.01 M phosphate buffer (pH 7.2) for at least 24 hours at 4°C with constant stirring. Change the entire buffer volume at least once, preferably at 9-12 hours. Each group will require 2 mL of dialyzed serum. Part B. Preparation of DEAE cellulose 1. To prepare the diethylaminoethyl (DEAE) cellulose, follow the instructions in the Whatman DEAE manual precisely. From the old Manual: A. Dialysis of human serum. 1. Dialyze 30 mL of human serum against 0.01 M phosphate buffer (pH 7.2) for at least 24 hours at 4°C with constant stirring. Change the entire buffer volume at least once, preferably at 9-12 hours. Each group will require 2 mL of dialyzed serum.

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B. Preparation of DEAE cellulose. 1. Begin this procedure the day before lab. 2. Stir 25 g of DEAE cellulose into 500 mL of 1 M phosphate buffer (pH 7.2)(i.e., 20 mL of buffer/gram DEAE cellulose). Allow the slurry to settle for 10 min and discard the supernatant. 3. Add 500 mL more buffer to the DEAE cellulose, and again stir and allow to settle. Collect the supernatant and measure its pH. 4. Continue washing the DEAE cellulose until the supernatant has a pH of 7.2. This may require several changes of buffer. 5. When the pH reaches 7.2, wash the DEAE twice with 0.001 M phosphate buffer (pH 7.2). Store the matrix in 0.01 M phosphate buffer in the refrigerator until lab. Just before use, remove most of the buffer and mix the slurry thoroughly before use. 6. Add 10 mL of DEAE slurry to 15 mL centrifuge tubes. Prepare 1 tube for each student. Chapter 8. Protein Fractionation. SDS-PAGE. Apparatus Electrophoresis chambers Power supplies and leads Micropipettors Water bath set to boiling (or a beaker in the microwave) Rocker table Supplies IgG sample from last week Sample buffer Running buffer (10x tris/glucine/SDS from Bio-Rad, cat no. 161-0732) Coomassie blue stain (Bio-Safe Coomassie for Bio-Rad, cat. No. 161-0786) Microcentrifuge tubes Gel loading tips microfuge tubes gel loading tips zip-lock bags Sample buffer Prepare the sample buffer by adding 4.0 mL dH2O 0.5 M tris-HCL, pH 6.8 1.0 mL glycerol 0.8 mL 10% SDS 1.6 mL -mercaptoethanol 0.4 mL 1% bromphenol blue 0.2 mL Aliquot into 0.5 mL fractions and store at -20°C.

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Chapter 9. Protein Fractionation: Western Blotting Apparatus Rocker table Small plastic trays 6" rulers 25 mL graduated cylinder Supplies Blocking solution (KPL cat. no. 50-82-00), 25 mL/blot. Wash buffer (KPL cat. no. 50-63-00), 80 mL/blot. TMB membrane peroxidase substrate (KPL cat. no. 50-77-03) Goat anti-human IgG-HRP conjugate (BioRad cat. no. 172-1050). Dilute 1:1000 (20 L in 20 mL Blocking solution). Chapter 10. Supplementing RPMI 1640 Culture T27A cells in RPMI 1640 supplemented with penicillin, streptomycin, L-glutamine, 25 mM HEPES, and 10% bovine calf serum. 1) Check the bottle of RPMI 1640. It may already contain all the above supplements, except bovine calf serum. If so, add 50 mL BCS (Hyclone # SH30072.03) to a 500 mL bottle of RPMI 1640. Be sure to use sterile technique and work in the hood. 2) If the medium needs antibiotics and L- glutamine add 5 ml of L-glutamine / penicillin / streptomycin solution (Sigma # G-1146) to the 500 mL bottle. Chapter 11. Cell Culture: Primary Culture of Chick Embryo Fibroblasts. Procedure Apparatus CO2 incubator; 37°C, 5% CO2. Check CO2 level in bottle. inverted microscopes compound microscopes bench top centrifuges stir plates Bunsen burners and strikers electric pipette pumps hemocytometer and cover slip

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Material Chick embryo, 8-11 days old. Call Linda Moore at Hatchery #2 (410-543-3478) to arrange to pick up eggs. 70 % ethanol (v:v), (sterilizing agent) plastic forceps in 70% ethanol Petri dishes sterile Hanks' balanced salt solution (HBSS), 12 mL/pair plus enough for diluting for cell counts sterile 125 Erlenmeyer flask with sterile stir bar tissue culture flasks 2/group trypsin/versene 18 mL / group CEF culture medium 20 mL / group (portion out in 50 mL tubes) plus some filled tubes 50 mL centrifuge tubes 12 × 75 mm test tubes 0.04% trypan blue in PBS sterile pipettes, 1 mL and 10 mL CEF culture Medium. Mix the following: F-10 Medium 376 mL M-199 medium 376 mL Newborn calf serum 40 mL 100x antibiotic/mycotic 8 mL Chapters 12 and 13. Cell Culture: Secondary Cultures of Chick Embryo Fibroblasts. Apparatus CO2 incubator; 37°C, 5% CO2 inverted microscopes compound microscopes bench top centrifuges electric pipette pumps hemocytometer and cover slip Material 70 % ethanol (v:v) tissue culture flasks trypsin/versene CEF culture medium 15 mL centrifuge tubes 12 × 75 mm test tubes PBS 0.04% trypan blue in PBS sterile pipettes, 1 mL and 10 mL Ca2+ and Mg2+ -free phosphate buffered saline

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Appendix 2. Instructions to Authors from JCB.

An excerpt from the on-line Instructions to Authors from the Journal of Cell Biology (http://www.jcb.org/misc/ifora.shtml). Manuscript organization and preparation Conventions. The JCB follows the abbreviations and other conventions of Scientific Style and Format: The CBE Manual for Authors, Editors, and Publishers (6th Edition, 1994, Council of Biology Editors, Inc., 9650 Rockville Pike, Bethesda, MD 20814). For chemical nomenclature, follow the Subject Index of Chemical Abstracts. Capitalize trade names and give manufacturers' names. Gene names must be italicized. Authors must use the original name published for a gene unless they have obtained permission to rename the gene from the authors of the original study (or from a governing body such as, in the case of a yeast gene, the Saccharomyces Genome Database curator). American spelling should be used throughout the manuscript. Please use Symbol font for all Greek characters. Title page. The title should be <100 characters (not including spaces). Provide the complete names of the institutions where the work was done, and the name, mailing address, telephone number, fax number, and email address of the author to whom correspondence and proofs are to be sent. If you wish to have two corresponding authors listed for the paper, you must designate one of them to communicate with the editorial and production offices. If a change of address is imminent, indicate the change and the date effective. Furnish a condensed title of <45 characters for incorporation in the running head. A revised manuscript should include the original manuscript number and the word "Revision." The number of characters must be listed on the title page. Abstract. The abstract may be paragraphed and should give a synopsis of the work reported that is self-explanatory and suitable for use without changes by abstracting services. Abstracts must not exceed 160 words. References must be cited in full in the abstract. Abbreviations. A term that does not appear on the JCB standard abbreviations list must be used at least three times in a paper to qualify as an abbreviation. Spell out the term on first mention, and follow it with the abbreviated form in parentheses. Thereafter, use the abbreviated form. Supply a list of nonstandard abbreviations used in the paper, in alphabetical order, giving each abbreviation followed by its spelled-out version. Materials and methods. Please see the JCB policy on image acquisition and manipulation above. An 'Online Supplemental Material' section, providing a brief description of any materials submitted for online only publication (such as videos, data sets, or supplemental figures), must appear at the end of Materials and methods section. References. References should be cited parenthetically by author and year of publication. If automatic referencing systems are used, the references must be finalized and reduced to text before submission. References should be listed alphabetically by first author's last name. The authors must be cited in the order in which they first appeared in publication and as they

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subsequently appear in Medline, even in cases where more than one author contributed equally to the work. Include all authors' names (do not use "et al."), year, complete article title, and inclusive page numbers. See examples below. Abbreviate the names of journals according to PubMed; spell out the names of unlisted journals. Unpublished material should not appear in the reference list. Citations such as "manuscript in preparation," "manuscript submitted," "unpublished results," "unpublished observation," and "data not shown," must appear parenthetically in the text as "unpublished data." As an alternative to "unpublished data," additional images, video, data sets, or methods not shown in the article may be included as Online Supplemental Material. When a person(s) who is not an author of the article is the source of unpublished data, those data must be cited as a "personal communication." In the case of "personal communications," authors must provide a signed letter of permission from the source of the communication authorizing the authors to cite the communication. Unpublished work may not be cited in the Materials and methods section. Citation of abstracts in the reference list is not permitted; these should be incorporated parenthetically into the text, giving the authors' names, meeting name and year, and abstract number. Adhere to the reference formats provided by the following examples:

Journal Articles

Two authors: Yalow, R.S., and S.A. Berson. 1960. Immunoassay of endogenous plasma insulin in man. J. Clin. Invest. 39:1157-1175. More than two authors: Benditt, E.P., N. Ericksen, and R.H. Hanson. 1979. Amyloid protein SAA is an apoprotein of mouse plasma high density lipoprotein. Proc. Natl. Acad. Sci. USA. 76:4092-4096. In press: Brown, W., and A. Nelson. 1983. Phosphorus content of lipids. J. Lipid Res. In press.

Online Peer-Reviewed Articles

Published article with only DOI: Lopez-Soler, R.I., R.D. Moir, T.P. Spann, R. Stick, and R.D. Goldman. 2001. A role for nuclear lamins in nuclear envelope assembly. J. Cell Biol. doi:10.1083/jcb.200101025 Published article with both DOI and pagination: Lopez-Soler, R.I., R.D. Moir, T.P. Spann, R. Stick, and R.D. Goldman. 2001. A role for nuclear lamins in nuclear envelope assembly. J. Cell Biol. 154:61-71. doi:10.1083/jcb.200101025.

Complete Books

Myant, N.B. 1981. The Biology of Cholesterol and Related Steroids. Heinemann Medical Books, London. 882 pp.

Articles in Books

Innerarity, T.L., D.Y. Hui, and R.W. Mahley. 1982. Hepatic apoprotein E (remnant) receptor. In Lipoproteins and Coronary Atherosclerosis. G. Noseda, C. Fragiacomo, R. Fumagalli, and R. Paoletti, editors. Elsevier/North Holland, Amsterdam. 173-181.

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Tables. Files for tables should be included with your submission. Double space tables on pages separate from the text and make them self-contained and self-explanatory. Do not use vertical rules. Label each table at the top with a Roman numeral followed by the table title. Insert explanatory material and footnotes below the table. Designate footnotes using lowercase superscript letters (a, b, c) reading horizontally across the table. Supply units of measure at the heads of the columns.

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