Molecular and Cell Biology Lab

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In Cell and Molecular Biology, we are going to give you what is probably your first .... 11. Never start a sentence with a digit, always spell the number out if it is the first ... Try to condense each section of the paper (i.e., Introduction, Materials and ...... M38 cell response (record response of second cell line on separate sheet).

Molecular and Cell Biology Lab Manual – Dr. Menze BIO 3120 – Sec I (Spring 2013)





Guidelines for Writing a Scientific Paper and Course Dynamics

Chapter 1



Chapter 2.1 Restriction Digestions of DNA


Chapter 2.2 Electrophoretic Separation of DNA


Chapter 2.3 Isolation and Quantification of Plasmid DNA


Chapter 2.4 Restriction Mapping of Plasmids


Chapter 2.5 Ligation of DNA


Chapter 2.6 Transformation of E. coli


Chapter 3

Polymerase Chain Reaction


Chapter 4

Enzymatic Activity of Gene Products (Proteins)


Chapter 5.1 Extra Credit Lab - Cell Counting Contest


Chapter 5.2 Sterile Cell Culture: Designing an Experiment


Chapter 6

Western Blotting


Chapter 7

Gene Reporters


Chapter 8



Chapter 9

Molecular Adaptations to Temperature


Appendix 1 Lab Report Checklist.


Appendix 2 Bioinformatics 2


Appendix 3 Bioinformatics 3


Appendix 4 Overview of Eukaryotic Transcription/Translation


Appendix 5 Syllabus and Tentative Schedule



Prefix Writing a Scientific Paper A. Introduction In Cell and Molecular Biology, we are going to give you what is probably your first experience writing a paper in the style used by scientists. This is neither 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. B. Objectives This chapter provides both general and specific guidelines for preparing a manuscript for publication in a scientific journal. These are the guidelines you will follow to prepare your laboratory report for this course. Follow these carefully and refer to them often while preparing your report. Use them later in your career to write scientific manuscripts. Your laboratory report will be based on a series of experiments that was designed by you and your group. You will be graded based on your group effort and your final report. C. Instructions The best way to learn this style is to use it in writing, and to 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 report. You will also read examples that you will find in the current literature. In fact, your first assignment is to obtain (from the library or from an electronic source, see below) a journal article of your choice (any field or discipline of the biological sciences), to use as an example when you prepare your lab report. 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 substantially if you carefully refer to these guidelines and the examples before your prepare your report. Your lab report is based on a series of experiments that your group will design to characterize a genetically engineered (Kc167-LEA-GFP) and a wildtype cell line (Kcs167) from the fruit fly Drosophila melanogaster. You will need to read the outline of the experiment in advance (see page 56) and decide with your group on a specific experimental set-up. You will be graded based on your group effort (design of the experiment and hands-on work) and your written report. After

4 consulting with me you may decide to submit your manuscript (lab report) to your electronic writing portfolio. D. The Scientific Paper A scientific paper is usually an original article published in a scientific journal. It is a written description of new research results. In its pre-publication form it is called 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 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 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 (Lab Report) should strictly adhere to this format. The ultimate source of instructions on how to prepare a manuscript for a given journal is the “Instructions for Authors” or “Guide for Contributors” that can be found on the webpage 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.

5 E. Group Assignment 1 – Due the first week of laboratories: 15 points A. Examine the “before” and “after” examples of a scientific article on d2L. These represent “real life” examples of a paper that was sent to a journal and subsequently published (in this case in the Journal of Biological Chemistry). The “before” paper is called a manuscript and is what was mailed to the editor of the journal. The “after” version is what the publisher‟s type-setter produced in the publisher‟s office and what was actually published (after it was checked for correctness by the authors). 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 a basic manuscript based on your laboratory work that is precisely correct in format is a major goal of this course. Assignment 1: Write a one page summary about the FIRST sample article as a group. Compare and contrast the differences between the manuscript and the paper, and summarize the major findings of the article. You may want to consult the Addendum article as a source of background information. An Addendum article is a short advertisement that summarizes the major findings of the primary research publication. B: Find an article of interest to you from a respected, peer reviewed journal of your choice. This article will become your guide for writing your own reports. Be sure that the article you chose has all the sections mentioned above, i.e., Introduction, Methods and Materials, Results, Discussion, etc…. Bring this article to lab with you the second week (first week of lab) and keep it handy to use as a general guide throughout the semester. In case your group cannot decide on an article consult with me. C: Find the “Instructions for Authors” for the journal in which your example article was published. Bring your summary, journal article, and instructions for authors to the first laboratory section on Tuesday, 01/15. F. 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. Appendix 1 at the end of this Lab Manual is a checklist that allows you to easily check for most of these items quickly. Though there are few rules concerning scientific writing that can be “written in stone”, the following come close and should be kept in mind as you write.

6 1. Double space the entire document. 2. Write clearly and be brief. Scientific literature must be concise, terse, and succinct. 3. Avoid vernacular (e.g., slang) and jargon as much as possible. When you must use it, explain it on its first use, if possible. 4. 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 potassium chloride (KCl)…” and not “The concentration of KCl (potassium chloride) was…” 5. Avoid the use of phrases like "It can be seen that…”, “It is clear that…”, “We concluded…" and "We can see…” 6. Never refer to the Instructor of the class. Shun phrases like “The instructor prepared…”, “Dr. Menze gave us…” and “We obtained from the instructor…” You will never do this in a real article. 7. Verb tense may changes between sections; past tense is usually used for describing procedures (i.e., Materials and Methods) and results (Results), and present tense is used to describe conclusions (Discussion). However, be careful not to change tense within a section. 8. Include headings for each section of your paper. Follow the “Instructions for Authors” of your guide article, or the SECOND example article posted on d2l. 9. 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. 10. 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. 11. 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 ml of reagent A…..” rather than “5 ml reagent A…” 12. Carefully proof read your drafts to catch any errors and also to fine-tune content and flow. Finally, it is well known even by 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

7 you do it, the more you will appreciate this fact. Do not be frustrated or feel 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. F. Writing the Manuscript: The Components of a Scientific Paper I. 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. Good titles do not include phrases such as “Results of …”, "Investigations of ...", "A study of ...", "Observations on ..." 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 extract keywords that are used in cross-referencing and computer searching. An improperly titled paper may never reach the audience for which it was intended, so be specific and keep these facts in mind when preparing your title. On the line(s) just below the title should appear the names of all the authors (the members in your group). Since you are preparing the manuscript you are consider being the „first author‟ and you should put your name first. Below that appear the author‟s affiliations and other contact information. Follow the sample manuscript on d2L. Give your report a Title Page consistent with the guidelines presented above. However, do not give your real phone number and home address! Use the Department of Biological Sciences as your address for correspondence and make up a phone number. II. Abstract Write the abstract last so that it accurately reflects the content of the paper. The abstract should briefly state the issue or problem at hand, the major objectives of the work, the methods used (in a sentence or two and avoid details), and a summary of the overall conclusions. It is not necessary to repeat information that appears in the title. Try to condense each section of the paper (i.e., Introduction, Materials and Methods, Results, and Discussion) into one or two sentences each. A good abstract will allow a reader to identify the basic content of the article and give them an idea of the value of reading the entire study. Make it accurate and interesting! Limit the size of the Abstract to 250 words. The title and abstract must constitute a “stand alone” document. They are published separately from the article itself in abstracting and citation services like PubMed and Web of Science. Try to leave out references (though

8 you will often see these in abstracts) and references to tables or figures. Spell out, or better do not use, abbreviations and acronyms even if they are defined in the body of the article. 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: 1. 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 or conclusions). 2. Do not include literature citations in your abstract. 3. Do not include acronyms, spell everything out. III. Introduction As with the manuscript, the Introduction does not have to be long to be good. Introductions are generally one to five paragraphs long, but can be longer if you believe it to be necessary to get your points across to your readers. It should clearly state the hypothesis, the background that explains the hypothesis, and the reasons for doing the study. Also state how your work differs from published work or how it is related to those works. The Introduction should begin with an explanation of the issue or problem at hand and a brief evaluation of the pertinent literature (the lab report must have a minimum of ten references). It is not necessary to try to impress the reader with the quantity of material you found on the topic. Always use prose and never use a “cookbook” format or a series of point-wise statements to make your arguments. Demonstrate an understanding of the relevant and important issues or problems in the field and point out how the work in this article adds to the overall body of knowledge on the subject. Clearly state the objective of the work. The Introduction can end with either statement of objectives or a summary of the major findings of the work. In the Introduction it is essential to give the reader an idea of where the paper is heading and what to expect in the pages ahead so that they can follow the evidence you are about to present. The main goal of the Introduction is to set up your reader for what is to follow. Tell them what is about to follow, why it is important, what you are going to do, and how you are going to do it. Your Introduction should comprise one to four paragraphs. Try to accomplish the objectives listed above in about two pages of text. Your references will occur primarily in the Introduction and Discussion sections. The lab report must include a minimum of 10 primary and secondary references. NO URL’s (In a primary reference, the authors conducted the experiments that are described in the

9 article. You should cite mostly primary references). You will find suggested references on D2l. IV. Materials and Methods This section is also called "Experimental Methods" in some journals. In this section you explain to your reader what you did to collect the data that are going to be presented in the Results section. Your goal in this section of the paper is to provide enough detail of what you did and how you did it so that other competent scientists will be able to repeat your experiments and reproduce your results. If your results are consequential, they will be reproducible. As a scientist, it is your responsibility to provide the information necessary for others to repeat your work. Always write the M&M section as a past tense description of the experiments you performed. Again, it is important to use prose and not to write a “cookbook” description of your methods. This section is often divided by sub-headings, for example: “Cell Culture”, “Determination of cell Viability", "Chemicals", "Statistics", etc… Equipment and materials available directly from commercial suppliers should be described exactly (e.g., Hauser, hematocytometer) and the source should be provided. For example, “…potassium chloride was obtained from Sigma Chemical Company (St. Louis, MO).” The methods used to set up equipment and to prepare reagents should be stated precisely. In this regard is it common to refer to apparatus and recipes described in the literature, e.g., “Kc167 cells were cultured as described by Cherbas (Cherbas et al., 1999). It is unnecessary to include descriptions of procedures that are familiar to most scientists. For example, it is not necessary to say “We added 1 g of KCl to 95 ml of distilled water, and then brought the mixture up to 100 ml”. Instead say “We prepared a X M solution of KCl” or just “ X M KCl 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 “cells were seeded into well number 1,” or “six wells were labeled 1 through 6”. Do not include tube numbers or the “cookbook” tables found in your lab manual; instead describe the procedures you used. 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 carried out. Be precise in describing measurements and be sure to use standard deviation (SD). Ordinary statistical methods can be used without comment (e.g. Students t-test; ANOVA), but advanced or unusual methods should be accompanied by a literature citation. It may also be appropriate to explain why you chose to measure the variables you are reporting and where relevant, state the conditions prevailing when measurements were made if they are not implicit in the experimental design (e.g., pH, temperature, humidity, time of day, etc...).

10 Follow the directions above. This section should be as short as practical. Do not be worried that this section is “too short”. Short is good. Short is what you want. Make sure a competent researcher could duplicate what you did, and be as brief as possible doing it. Follow the examples you have available. V. Results Just like it states, in the Results section you present your results. Present the data and point out trends and other pertinent observations. Present your figures and tables here, as well as your statistical analyses. Remain objective and present a logical flow of data and concepts. Your goal is to take the reader on a guided tour of what you found after doing the experiments described above in the M&M section. Convince the reader that your arguments are both rational and valid. Any lack of clarity may lead to doubt. If your readers are confused or unable to follow your logic, they may not recognize the significance of your work, or worse, they may reject your ideas completely. Before you even start writing, make sure you understand your data! Are they clear to you? Are they telling you a story and do you understand it? Only after you have a thorough understanding of your data should you start to plan the best ways to present them to a reader. If a result is simple, state it in the text. More complicated results will require you to present tables and figures. The decision of whether to use a table or a figure to present a given set of data is yours. If the exact numeric values of the data are important then use a table. If patterns or trends are what you want to convey, use a figure. Sometimes this decision is a difficult one. Experience with this will help you. Tables and figures with their titles and legends (respectively) should be able to stand alone without reference to the text (but the placement of titles and legends are different for table and figures, see below). The Results section should be short and sweet. Do not babble. Be terse. For example, do not say "It is clear from Fig. 1 that cell growth increased with the amount of serum added to the medium". Say instead "Cell growth increased with added serum (Fig. 1)". Avoid presenting long lists of results with no accompanying explanation or analysis. "Serum level significantly affected growth (Figure 1). pH significantly affected growth (Table 1). Light levels had no significant effect on cell growth (Figure 2)." Develop each idea and describe the effect. In what ways did the independent variable change? The Results section should only include direct interpretations (“the cells grew”, “the enzyme was active”, “the surviving cells expressed LEA-GFP”). However, be careful! In the Results section you do not include any speculation or discussion of the observations you make. This is one of the harder skills to master when writing a scientific article. Speculation and discussions belong in the Discussion section. Indirect interpretations (“the expressed LEA protein may help these cells survive when…”) also belong in the Discussion section. DO NOT FOLLOW THE EXAMPLE 1 ARTICLE! This article is

11 considered a „short communication‟ and combines Results and Discussion into one section. Your manuscript will contain a separate Result and Discussion section as in EXAMPLE 2. By convention the first table presented in the Results section is “Table 1” and the first figure referred to, even if it is after table 1, is called “Figure 1”. It is a good idea to put off numbering your tables and figures until 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. Also, once again, brevity is desirable. Be as brief as you can while still giving your reader everything they need to know to follow your “story”. I 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. You may only need a few sentences or you may need several paragraphs. Keep it minimal. SHORT, SHORT, SHORT! No fluff! VI. Preparation of Tables and Figures A. General Considerations: 1. Present data in either a figure, a table, or in the text, but never present the same data twice. 2. In the text of the paper always refer to every table and figure included. 3. Do not present data in a table or figure when it could easily be included in a sentence or two of text or incorporated into the title of a table or figure legend. 4. Do not title figures, but be sure to give them legends. 5. Present figure legends together on a separate page. Figure legends should not be printed on the Figure itself. 6. Each figure should appear on its own page. See the EXAMPLE 2 manuscript on D2l. 7. Numbers that are presented in the columns of tables and along the axes of figures should always have the same precision (i.e., the same number of decimal places). Also, numbers should never start with a decimal point; always include a leading zero (e.g., 0.1, never .1). 8. Never start a figure or table legend with “A graph showing…”, “A Figure of…” or “A Table showing….”. Refer to the examples you have available.

12 B. Tables: 1. Tables, should have at their head (i.e., at the top) the table number followed by a legend. The legend should be brief, but should allow the table to stand alone. The reader should be able to understand it without referring to the text. See the examples on D2l and in the sample article you got from the library. 2. Include appropriate and precise column headings. 3. Do not include columns of data that have the same value repeated in every cell. If the repeated value is important to the table put it in the title. C. Figures 1. Always include, on a separate page, the legends (some journals call them captions) describing each figure. Legends must be brief, but remember the figure and its legend must be able to stand alone without referring to the text. 2. Label both axes with a brief but informative title. Always include units of measurement on the axes! 3. A figure should not fill the entire page. Leave ample room for axis numeration and titles. 4. Match the extent of the axes to the data. For example, if the data range between 0 and 190, the axis should extend to 200, or maybe 250, but not to 500 or 5000. 5. Each figure should be placed on a separate page (but they do not need to cover the entire page!). 6. Do not place figure legends on the graph itself. Identify symbols in the text of the figure legend. 7. 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. Meticulously follow the directions given above. Be sure to re-read these instructions and examine the examples you have before beginning to prepare you Results section. VII. Discussion The Discussion is where you explain and interpret your results. Relate your explanations and interpretations to other work in the field. Conclusions are presented and discussed in this section. Base your conclusions on the evidence you presented in

13 the Results section. Explain how your goals and objectives (stated in the Introduction) were met. Was your hypothesis supported or countered? (E.g. Does LEA-GFP increases osmotic stress tolerance of Kc167 cells?) Explain the significance of results and how they add to what was previously known. How do your findings relate to those of others? Have new biological principles been established, confirmed or reinforced? What generalizations can be made? Are your results the expected ones or were you surprised? What are the practical implications of your results? What are the theoretical implications of your results? As you discuss what you found always remain firmly within the evidence you presented. Be careful not to overstep and draw conclusions that are not directly supported by your results. You may choose to speculate, but do not let it form the bulk of the discussion. You can make suggestions for future research. You can explain any methodological errors made during the course of the experiment. However, NEVER say “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, do not leave the reader thinking "So what?” End the Discussion with a short summary of what was found and why it is important. For our purposes, aim for a Discussion one page to two pages in length. It is important to include references in this section, and all lab reports must have a minimum of ten references. Sometimes journal allow you to publish shorter articles in which the Results and the Discussion part are combined into „Results & Discussion‟ (see EXAMPEL 1 on D2l). Since this is rather an exception you will need to write your lab report with separate Results and Discussion sections. VIII. References Whenever you use information obtained form a source other than your own head, you must acknowledge that source. Other, non-scientific forms of writing rely greatly on the use of quotations. In scientific writing quotes are almost never used (I don‟t know why, but it is so). In scientific writing information from published sources is extensively reworded and immediately followed by an indication of the source. The list of these references is then included at the end of the paper. 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 Experimental Biology (the Instructions to Authors for JCB can be found at: (Use this format regardless of the “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.) Each reference cited in the text must be listed in the References and vice versa: please check these carefully.

14 Literature citations in text are as follows. (1) One author – (Jones, 1995) or (Jones, 1995; Smith, 1996). (2) Two authors – (Jones and Kane, 1994) or (Jones and Kane, 1994; Smith, 1996). (3) More than two authors – (Jones et al., 1995) or (Jones et al., 1995a; Jones et al., 1995b; Smith et al., 1994; Smith et al., 1995). (4) Avoid any additional text within the brackets; this format is necessary for on-line literature searches. (5) Manuscripts accepted for publication but not yet published – list in References as (in press). (6) Citation of unpublished work: (a) Your own unpublished observations and results submitted for publication should be cited in text only and not in the reference list. Use the format (S. P. Jones, unpublished). (b) Personal communications, i.e. the unpublished observations of other scientists, will only be published when substantiated by written permission. Reference list (1) References are listed in alphabetical order according to surname and initials of first author. Within a group of papers with the same first author, list single author papers first, then papers with two authors, then et al. papers. If more than one reference exists for each type, arrange in date order. Use a and b for papers published in the same year. (2) 'In press' citations must have been accepted for publication and the name of the journal or publisher included. (3) Initials should follow all surnames in the list of authors; insert a full stop and space after each initial and parentheses round the date followed by a full stop. Use bold for authors' names. (4) Use USA National Standard abbreviations for journals. (5) Use the following style: Rochlin, M. W., Itoh, K., Adelstein, R. S. and Bridgman, P. C. (1995). Localization of myosin IIA and B isoforms in cultured neurons. J. Cell Sci.108, 3661-3670. Matlin, K. S. and Caplan, M. J. (1992). Epithelial cell structure and polarity. In The Kidney: Physiology and Pathophysiology (ed. D. W. Seldin and G. Giebisch), pp. 447473. New York: Raven Press Ltd. If there are more than 10 authors, use 'et al.' after the 10th author.


Chapter 1 Handling Micropipettes A. Introduction In many cases, the molecules that scientists will want to manipulate exist at very low concentrations in living systems, such as the sub-micromolar ranges. In order to effectively carry out our experiment, it is essential that we be able to accurately deliver, extract, and transfer quantities of liquid that are minute. The invention of micropipettes by Dr. Gilson and Dr. Lardy in 1974, with the subsequent generations of modifications and improvements, has led to the micropipettes that are both ubiquitous and indispensable in modern biology laboratories.

The Power of 10 10


SI unit




















All the modern micropipettes nowadays operate on the same principle of air displacement. In general, the user adjusts the micropipette so that a fixed amount of air is expelled from the barrel. Then, holding this negative pressure, liquid is suctioned up into a temporary holding tip, and then subsequently delivered to the desired place. In other words, the micropipettes operate on the same principle as a straw. B. Objectives In this exercise, students will learn how to operate the micropipette to achieve accurate delivery of liquid reagents.

C. Terms Micropipette

A mechanical device capable of allowing the delivery of minute quantities of liquid

Pipette tips

Plastic, disposable tips that fit onto appropriate micropipettes

Milliliter (ml)

A unit for the volume of liquid, 10-3 liter

Microliter (µl)

A unit for the volume of liquid, 10-6 liter

Microcentrifuge tubes

Plastic, disposable tubes that hold 0.5 ml, 1.5 ml or 2.0 ml volume. Commonly used in molecular biology applications.

16 D. Background There are two important steps to accurate micropipetting: 1. Choosing the right micropipette 2. Correct operation of the micropipette 1. Choosing the right micropipette In this class, students will be exposed to 4 different micropipettes. Each one has a designated range of liquid that it is designed to handle. It is critical that the volume to be manipulated falls within the stated range of the micropipette to ensure accuracy. For example, if we need to transfer 20 µl of liquid, we cannot use the 0.2-10 µl micropipette, and we should not use the 100-1000 µl micropipette. Instead, we should choose the 10-100 µl micropipette. 2. Correct operation of the micropipette 

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Micropipettes are precision equipments that require high levels of calibration. Misuses of micropipettes not only result in inaccurate results for your experiment, but in some cases can actually damage the equipment so that it will not be accurate in future uses. The most common error is the attempt to adjust the volume of the barrel beyond its stated range. NEVER ATTEMPT TO SET THE VOLUME OF A MICROPIPETTE TO BEYOND ITS STATED RANGE. The volume to be delivered should be set on the micropipette prior to the operation. Use the volume adjustment knob to set the correct volume and then lock it to prevent accidental changes of the setting during manipulation. Before use, the micropipettes must be fitted with the appropriate tip. The three kinds of tips are white (0.2-10 µl), yellow (10-100 µl), and blue (1001000 µl), corresponding to the color-coded micropipettes. To fit a tip onto the micropipette: insert the micropipette into the tip, and then tap twice firmly. The tip should be snug and stable on the micropipette. The plunger of the micropipette has three positions: rest, first stop, and second stop. As the plunger travels from the rest to the first stop, it will expel air equals to the volume set on the micropipette. The user should hold the plunger in that position as s/he inserts the micropipette tip into the liquid. Make sure that you are holding the micropipette vertically, and that you place the tip into the sample to the proper depth (1 – 2 mm for 10 µl, 2 – 3 mm for 100 µl, 3 – 6 mm for 1000 µl). Make also sure that the tip will remain in the liquid after the set volume is removed. Then, while the tip is fully submerged, gently release the plunger as it springs back to the rest position. At this point, the tip holds a volume of the liquid equal to the volume set on the display. To deliver the liquid, insert the tip into the new container, and slowly depress the plunger to the first stop. Then, press further into the second stop, and hold the plunger at the second stop while moving the tip away. Once the tip is away from the liquid, release the plunger back to the rest position.

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Since the target volume is small, the liquid is susceptible to static electricity and surface tension factors. This means that sometimes, the liquid reagent may become stuck to the outside of the tip, instead of being delivered into the tube. To avoid this, make sure that the tip is submerged into the second liquid while releasing. If the target is an empty tube with no liquid, touch the micropipette tip to the bottom of the empty microcentrifuge tube while releasing. To avoid cross-contamination of reagents, tips should be refreshed after each use. To remove the tip, hold the micropipette over the waste receptacle provided and push the eject button.

Figure 1.1. The correct way of using a micropipette.

3. Summary: A. Choose the micropipette with the correct range. B. Adjust the volume of micropipette and lock it. C. Fit micropipette tips to the micropipette. D. Depress plunger to first stop, hold. 1. Submerge micropipette tip into liquid. 2. Gently release the plunger back to rest position. 3. Move micropipette tip out of the liquid and touch the container (e.g. micro tube) you like to transfer the liquid into with the pipette tip. 4. Gently depress the plunger to the first position, hold. 5. Push the plunger further to the second position, hold. Move the micropipette tip out of the container. 6. Release the plunger back to rest position and eject used tip.

18 E. Chapter 1

Each Student

Set up six micro-centrifuge tubes, and label them A through F. By using the three solutions provided (#1, #2, and #3), perform the following mixes. Table 1.1. Mixing chart Tube A B C D E F

Solution #1 4 µl 4 µl 10 µl 100 µl 80 µl 300 µl

Solution #2 4 µl 1 µl 3 µl 200 µl 0 µl 100 µl

Solution #3 4 µl 0 µl 1 µl 75 µl 45 µl 550 µl

Reminder: Bring your summary, journal article, and instructions for authors to this laboratory section. You will lose 15 points in case you didn’t complete your group assignment (see page 5)!


Chapter 2.1 Restriction Digestions of DNA A. Introduction Since the beginning of the last century, scientists have been interested in genes. New ways to study the structure, behavior, and activity of genes in more detail started to develop in the early 1970s. The ability to manipulate DNA to generate artificial recombinant forms of DNA is now a critical technique for the study of molecular biology. The ability to manufacture new combinations of DNA allows scientists to conduct experiments that address some of the fundamental questions regarding genetic control of living organisms. Broadly, the process of cloning genes into a new context is composed of 3 different techniques: restriction digestions of DNA, ligation of DNA, and the use of plasmids to transform new organisms. Since its discovery, gene cloning has become a routine and indispensable part of modern molecular biology. Before 1970 there was no method for cleaving DNA at discrete points. The breakthrough came with the discovery of an enzyme in Hemophilus influenzae that recognizes a particular target sequence in a duplex DNA molecule and breaks the polynucleotide chain within that sequence to give rise to discrete fragments. Restriction digestions are carried out by these enzymes and are called restriction endonucleases. These are produced by prokaryotes, and they have the ability to recognize specific DNA sequences and cleave the double-stranded DNA at those sites. Restriction enzyme systems allow bacteria to monitor the origin of incoming DNA and to destroy it if it is recognized as foreign. Many restriction enzymes exist, and there are many variations on their recognition sequences. Some enzymes recognize 4 bases, others, 12 bases. Some recognize perfect palindromes, while others do not. Some cut the DNA strands evenly, and others do not. The choice of which restriction endonuclease to use depends on the goal and the limitations imposed by the piece of DNA itself. In addition to gene cloning, restriction digestions are also used for other applications, such as Southern Blot and restriction fragment length polymorphism (RFLP). B. Objectives In this exercise, students will perform restriction digestions and analyze the results.

C. Terms Restriction endonuclease

An enzyme that breaks the double-stranded DNA at specific recognition sites

Recognition sites

DNA sequence that is recognized by restriction endonucleases


A perfect repeat from back to front, e.g. “Step on no pets.”


Palindromic sites

A stretch of DNA where reading 5‟ to 3‟ gives the same sequence in both directions, e.g. 5‟-CCATGG-3‟

Restriction buffer

A chemical solution, typically at 10X concentration, that contains the ions and other chemicals needed to enable activity of restriction endonucleases.

Recombinant DNA

Any artificially created DNA molecule which brings together DNA sequences that are not usually found together in nature.

D. Background There are several parameters to consider in setting up a successful DNA digestion. 1. Right Mixture Each digestion should contain the DNA, the enzyme, the buffer, and sterile water, in the right ratio. Too much or too little of one component can either impede or completely prevent the reaction from happening. The buffer we are using is 10X concentrated, so you always use 1/10 volume of your final digestion volume. The amount of sterile water to use is the final desired volume, minus the volume of the buffer, the DNA, and the enzyme. The amount of enzyme to use should be appropriate for the amount of DNA to be digested – the more DNA you want to digest, the more enzyme you will need. 2. Even Mixture All the components in the digestion should be evenly mixed so that the reagents are uniformly distributed. Since the volume you are handling for some of the components are small, please make sure that all the components are added appropriately and mixed evenly. You may need to use the microcentrifuge to collect all the volume. 3. Right Conditions Restriction endonucleases are enzymes; this means that their activities are affected by external conditions such as temperature, pH, and ionic strength. If the temperature is too high, it may denature the enzyme. If the temperature is too low, it will slow down its activities. The optimal temperature for each enzyme is given by the provider. The pH, and ionic strength is determined by the 10x buffer used. Some restriction enzymes show maximal activities in different buffer systems. However, the two restriction enzymes we will be using in this exercise, EcoRI and HindIII, exhibit high activities in the same buffer system.

21 E. Procedure

Each Pair

Each pair of students will receive a DNA sample of the plasmid: pPGK15.3. The plasmid contains a large DNA fragment, the liver-specific α-antitrypsin gene (α1AT). The two reactions to be performed will be: Table 2.1. Digestion of plasmid pPGK15.3

Component DNA EcoRI HindIII Buffer (10x) Sterile Water Final Volume 1. 2. 3. 4.

#1 (double digestion) 1 µg 1 µl 1 µl

#2 (control) 0.5 µg 0 µl 0 µl

50 µl

50 µl

Set up the digestions following table 2.1 Incubate your reactions at 37 oC for 30 minutes. Label sample CLEARLY!!! Store sample at -20 oC

This digestion will cleave the circular plasmid containing a mammalian α1AT gene into multiple linear fragments. After the restriction digestion is performed your products will be stored at -20 ºC. You will use the digested plasmid again in Chapter 2.2, 2.5, and 3.1. In Chapter 2.2 you will analyze the result of the restriction digestion via gel electrophoresis. In Chapter 2.5, you will create new plasmids in processes called ligation and transformation. In Chapter 3.1 you will see the impact of the digestion on PCR amplification of the a1AT gene.





HindIII PstI SalI AccI HincII XbaI BamHI AVaI SmaI SacI EcoRI


Fig. 2.1: (A) The plasmid pPGK15.3. (B) Principle of restriction enzyme digestion


Chapter 2.2 Electrophoretic Separation of DNA A. Introduction In many applications, it is important to be able to distinguish DNA fragments of different sizes. For example, perhaps only one of the two products from digesting a piece of DNA is needed, and we need to tell them apart by size. Or, perhaps, we are trying to determine the identity of the DNA using the sizes, such as during the forensic examinations of DNA. The most routine way of separating DNA by size is the use of agarose gel electrophoresis. In this approach, we capitalize on the over-all negative charge of DNA fragments and apply an electric current to mobilize them. Gel electrophoresis units are almost always simple electric circuits that can be understood using two simple equations. Ohm‟s law, V = IR, states that electric Figure 2.2.1. Basic setup of a gel field, V (measured in volts), is proportional electrophoresis apparatus. to the current, I (measured in milliamps) times resistance R (measured in ohms). Usually the gel itself provides nearly all of the resistance in the circuit, and the voltage applied to the gel will be essentially the same as the voltage applied to the circuit. The voltage gradient across the gel influences the electrophoretic mobility of the sample. An agarose gel is a complex network of polymeric molecules whose average pore size depends on the buffer composition and the type and concentration of agarose used. As these DNA fragments travel through the agarose gel matrix, larger pieces are impeded more than smaller pieces. P = I2R, states that the power produced by the system P (measured in watts), is proportional to the resistance times the square of the current. It is very important to know how much power a particular gel apparatus can dissipate and to monitor the temperature of gels run above that level carefully. Pure, solubilized DNA is invisible to the human eyes. Therefore, after the DNA has been separated during electrophoresis, the DNA must be visualized. To allow us to see the results of the DNA separation, stains are used. In this class, we use a chemical called GelRed, which will give off fluorescent light under ultraviolet excitation. Other chemicals commonly used to stain DNA include ethidium bromide (EtBr) and SYBR Green. Ethidium bromide was traditionally used for DNA staining; however, concerns about its possible mutagenicity and toxicity reduced its use. All these dyes exhibit a preferential affinity for DNA and the fluorescent signal is greatly enhanced when bound to DNA.

23 B. Objectives In this exercise, students will cast an agarose gel and perform electrophoresis to separate the digested fragments from Chapter 2.1. C. Terms Agarose Gel

Agarose powder that has been melted in the running buffer and then formed into a gel.


Small plastic pieces inserted into the agarose gel before it solidifies to create wells to hold DNA samples.

Loading Dye

A mixture of glycerol and tracking dyes. The glycerol weighs down the DNA sample by increasing its density so that it sinks to the bottom of the well instead of floating away. The tracking dye allows us to visually gauge the progress of electrophoresis.

DNA Stain

A chemical compound that can adhere to DNA and gives off fluorescent light when excited by UV light. Used to visualize DNA samples in agarose gels.

Running Buffer

A chemical solution (Tris-Boric Acid-EDTA, or, TBE, in this case) that contains the right ions to conduct electricity.

DNA Ladder

A pre-made mixture of DNA fragments of known sizes. It enables us to estimate the size of our DNA samples. In this class, we use the 100 basepair (bp) and the 1 kilobase (kb) ladders.

D. Background Briefly, agarose gel electrophoresis involves: 1. Melting of agarose powder to make the molten gel 2. Casting the molten agar to form a gel and inserting it into the electrophoresis apparatus 3. Filling the electrophoresis apparatus with the running buffer 4. Loading the DNA sample into the wells via micropipettes 5. Applying a defined electric current 6. Visualizing the results under UV light 1. Casting the gel It is important to make the molten agarose in the same solution as the running buffer to ensure uniform conductivity during electrophoresis. If the gel is made with a different buffer, the difference in conductivity will distort the separation and lead to inaccurate

24 results. Another important consideration is to allow the gel sufficient time to solidify. Gels that are still liquid in the middle will not give correct results. 2. Loading the samples DNA samples must be mixed with a loading dye to help it settle to the bottom of the well. The dyes migrate at a known speed, which helps us track the progress without having to take the gel out of the apparatus to look at it every time. For instance, the orange dye co-migrates with DNA at approximately 50 bp, whereas the blue dye comigrates with DNA at approximately 3 kb. To successfully load the sample, the tip of the micropipette should be either in the well, or just above it. Do not puncture the bottom of the well! Beginners may find it useful to use both hands and to rest their elbows on the bench while loading to help steady the micropipette. It is IMPORTANT to not move the gel apparatus following the loading of your sample, since movement of the tray will lead to the sample floating out of the wells. So be sure to position the gel box correctly before you start to load the sample. 3. Applying an electrical current The current we use to run the gels is lethal to humans. AT NO TIME SHOULD YOU TOUCH THE RUNNING BUFFER WHILE THE CURRENT IS ACTIVE. IF A GEL BOX IS LEAKING, DISCONNECT THE CURRENT FIRST. Safety features built into the apparatus will help reduce that risk, but do not be careless. 4. Visualizing the results The stain used to help us visualize the DNA requires activation by ultraviolet (UV) light. The gel, once finished, will be placed on the UV transilluminator and the resulting fluorescent image will be captured by a digital camera. Since UV light can cause damage to the human retina, students must avoid looking at the transillunimator when it is in use or wear protective eyeware. E. Procedure

Each Group/ Each Pair

Part I: Casting a gel (Each Group) 1. Fit the two rubber end pieces onto the gel tray and then rest the tray on a level surface. 2. Insert the comb at the right position – be sure that the teeth on the combs are not touching the rubber end piece and that they are approximately 5 mm away from the bottom of the tray. 3. Pour the molten agarose into the tray until it comes up about two-third of the way up the teeth of the comb. 4. Allow it to set for 20-30 min at room temperature. 5. After gel is set, carefully remove the comb and the stoppers. Be sure not to tear the gel.

25 6. Fit the tray into the box, making sure that the side with the well is near the negative end (–ve, cathode) of the box. 7. Add running buffer (1X TBE) to the box until both reservoirs are filled and the buffer completely covers the surface of the gel. Part II: Loading samples (Each Pair) 1. From each of your two tubes of products, remove 10 µl and spot it onto a piece of parafilm. 2. To each of your samples, add 2 µl of 6X loading dye. 3. Set the micropipette to 12 µl and gently pipette up and down to mix the dye with the sample. 4. Add each of your samples into a different well. Write down the sequence so you can interpret the results. 5. There should be 1 lane on the gel dedicated to each of the DNA ladders (serve as molecular weight standard). 6. Close the lid, check to make sure that the wells are closer to the –ve end. 7. Set the power supply current level to low, the display to mV, and set to 200 mV. Check to see that bubbles are coming out from the two ends – this will tell you that the unit is properly connected.

Fig. 2.2.2. Ethydium bromide stained DNA ladder (weight marker) on agarose gel visualized by UV light (1 kb DNA ladder, New England Biolab Corporation).


Chapter 2.3 Isolation and Quantification of Plasmid DNA A. Introduction Plasmids are widely used cloning vehicles which are replicons that are stably inherited in an extrachromosomal state. Most plasmids are double-stranded circular DNA molecules. Plasmids are widely distributed throughout the prokaryotes and vary in size from less than 1 x 106 daltons to greater than 200 x 106 daltons. Plasmids encode only a few of the proteins required for their own replication and in many cases encode only one of them. The use of plasmids is essential to gene cloning techniques. A large variety of plasmids are Figure 2.3.1. Structure of a vector commercially available, and most researchers (pCl-neo, Promega Corporation). adapt and modify these plasmids to suit their specific needs. The ability to obtain and manipulate plasmids is a standard technique expected of most practitioners of molecular biology. An ideal cloning vector has the following properties: a) low molecular weight, b) ability to confer readily selectable phenotypic traits to host cells, c) a single cloning site for a large number of restriction endonucleases. Quantification of DNA is an essential step in accurate gene cloning applications. The most commonly used method of DNA quantification is the ultraviolet light spectrophotometry (UV-Spec). In this method, the quantity and purity of the DNA sample is measured using the absorbance of the sample in 260 nm and 280 nm wavelength under excitation by UV light. The quantity is calculated using the 260 nm reading, whereas the purity is indicated by the 260/280 ratio. B. Objectives In this exercise, students will isolate plasmids from two different bacteria cultures and perform quantification and plasmid mapping on the product. Each bacterial strain contains a different plasmid and the students have to deduce which plasmid they isolated. C. Terms Resuspension

Solubilizing a solid in a buffer.


An insoluble solid left after centrifugation.


The liquid left after centrifugation.

27 D. Background 1. Cell Suspension Bacterial cells to be lysed are first collected and then the growth medium is removed. The bacterial cells are then resuspended in a buffer called GTE, which stands for Glucose-Tris-EDTA. The glucose is added to help maintain an osmotic pressure for the cells, so they do not lyse before we are ready. Tris is a buffer that works well to maintain pH 8.0 as a suitable environment for the DNA once the cells are lysed. EDTA is a common sequestrant that serves to chelate divalent cations especially magnesium. The enzyme that destroys DNA, called DNAse, requires Mg2+ to function. When EDTA is present, it chelates the Mg2+ ion and thereby inactivates DNAse, so that our plasmid DNA will not be degraded. 2. Cell Lysis The alkaline lysis procedure is the most commonly used method for isolating plasmid DNA from bacterial cultures. The SDS/NaOH solution contains sodium dodecyl sulfate and sodium hydroxide. SDS denatures bacterial proteins and NaOH denatures chromosomal and plasmid DNA. SDS is an anionic organic detergent and here it serves to dissolve the lipid bilayer of the cell membrane, so that our plasmid DNA can escape. The NaOH here is used at a concentration that will cause osmotic damage to the cells. This damages the cell walls and the leaky cell wall will now allow plasmid DNA to leave the cell and come into the solution. While there are many ways to lyse a bacterial culture, this is the optimal protocol because it does not destroy the entire cell, only makes it “leaky.” If the entire cell is disrupted fully, then the chromosomal DNA will also be precipitated in the following steps, thus contaminating our plasmid preparation with genomic DNA. 3. Potassium Acetate The addition of potassium acetate serves two functions: it neutralizes the pH after the addition of NaOH in the previous step, and the acetate will form insoluble complex with the dissolved lipid. This allows the cell debris to be separated out from the dissolved DNA in the centrifugation step. The mixture is neutralized with potassium acetate. This causes the covalently closed plasmid DNA to reanneal rapidly. Most of the chromosomal DNA and bacterial proteins precipitate as does the SDS, which forms a complex with potassium. The proteins and the chromosomal DNA are removed by centrifugation. However, make sure not to break the chromosomal DNA by excessive shaking (e.g. use of the vortex-mixer), which would result in poor separation of chromosomal from plasmid DNA. 4. Isopropanol The addition of isopropanol will turn the DNA into an insoluble solid. DNA is negatively charged due to its phosphate backbone. Isopropanol is much less polar than water; this

28 means that if enough alcohol is added electrostatic attraction between phosphate groups and the positive potassium ions present in solution become strong enough to form stable ionic bonds and precipitate DNA. This means that in presence of relatively high concentrations of monovalent cations isopropanol induces a structural transition in nucleic acid molecules which causes them to aggregate and precipitate from solution. At this step, it is critical that the isopropanol is evenly mixed with the solution, so that a uniform distribution of isopropanol is achieved. Uneven mixing will result in drastically lower yield of your final product. It is not uncommon to not see any solid at this stage, even though the DNA is precipitating out of solution. 5. 95% Ethanol Maximum DNA

Maximum BSA dsDNA Protein (BSA)


The use of 95% ethanol to wash the solid DNA will remove any residual salt and enhance your downstream application. However, ethanol itself can be an inhibitor of many reactions. Therefore, it is critical that you remove as much of the ethanol as possible, and then allow the pelleted DNA to dry thoroughly, before resuspending the material.


6. DNA Quantification 260

 280




It is useful, and sometimes essential, for molecular Wavelengtht, nm procedures to quantify the amount of DNA. The Figure 2.3.2. Absorbance spectrum unfortunate side effect of quantifying your sample of dsDNA and the protein bovine is that the amount of sample you used in the serum albumin (BSA). process can often not be recovered. Therefore, we will use only a portion of your sample, and we will obtain the true concentration by multiplying the result with the dilution factor. Based on the knowledge that 50 µg/ml of double stranded DNA (dsDNA) show an OD value (value of the A260 reading) of 1.0 at a light path length of 1 cm we can calculate the concentration of DNA in our sample. In the UV-Spec reading, you will see two readings for two different wavelengths: A260, and A280. DNA shows maximal light absorbance at λ= 260 nm. The A260 reading you obtain will be used to calculate the concentration of DNA in the tested sample. Concentration of original sample in g/ml = A260 reading * 50 * dilution factor Proteins show a local maximum in light absorbance at = 280 nm. You will use the absorbance at 260/280-ratio to determine the purity of your sample. Typically, a 260/280 ratio >1.8 indicates a sample that is relatively free of protein contamination. Values below 1.8 indicate that substantial amounts of proteins are present in your sample.

29 E. Procedure

Each Student

Part I: Plasmid Isolation 1. Transfer 2.0 ml of overnight bacterial culture A or B into a clean 2.0 ml microcentrifuge tube. Prepare two tubes per student. 2. Centrifuge at maximum speed for 3 minutes. 3. Remove the liquid using a micropipette. Dry blot the tube gently using a paper towel. Try not to disturb the pellet. 4. Add 50 l of ice-cold GTE buffer. Using the micropipette gently pipette the solution up and down to re-suspend the pellet. You want a completely homogenous solution at the end. Combine both tubes into one tube (You will have 100 l of combined pellets in the tube) 5. Add 200 l of SDS/NaOH solution. Mix by inverting the tube. DO NOT shake. Let sit at room temperature for 3 minutes. 6. Add 150 l of ice-cold KOAc. Mix by inverting the tube. DO NOT shake. Rest on ice for 3 minutes. 7. Centrifuge at maximum speed for 10 minutes. A large white solid mass should be visible on the side of the tube at the end. 8. Using a micropipette, transfer 400 l of the liquid into a fresh, new tube. Discard the tube with the white solid. 9. To the new tube, add 400 l of ice-cold isopropanol. Be sure to mix very well. Place at -20 ˚C for ~5 minutes. 10. Centrifuge at maximum speed for 10 minutes. A small white pellet should be visible on the bottom of the tube. 11. Using a micropipette, discard all the liquid. Be careful not to remove the pellet. 12. Add 200 l of 95% ethanol. Let it sit for 1 minute and centrifuge again for 3 min. 13. Remove all of the liquid, and let the pellet of DNA air-dry for 3 to 5 minutes. 14. Add 50 l of TE (Tris-EDTA) buffer to re-suspend the DNA.

Part II: UV-Spec Quantification 1. Let the TA load 4 l of the dissolved DNA to the micro plate. 2. The TA will take a UV-Spec reading of the sample and provide the results on D2l. 3. Calculate the concentration of plasmid DNA in your sample. Note your results and store the isolated plasmids at -20 ºC. You will need the plasmids AND your results generated for Chapter 2.4.


Chapter 2.4 Restriction Mapping of Plasmids A. Introduction If the sequence of a plasmid, or any piece of DNA, is known computer programs can be used to predict cutting sites based on the known restriction recognition sequences. Furthermore, programs are available that generate a “map” of the plasmid that features the relative distances of the restriction sites of interest. The distribution of cloning sites, or the plasmid map, is a key parameter for the utilization of plasmids because it determines which restriction sites are suitable for a particular cloning project. The generation of restricting maps of plasmids can also be done when the specific sequence of the plasmid is unknown. In this case, the specific distribution of cut sites can be seen as a sort of “finger print” for plasmids, so that examining the distribution of cut sites can often help us differentiate one plasmid from another. Both of these uses of plasmid maps are critical to gene cloning techniques. B. Objectives In this exercise, students will digest the plasmid they obtained in Chapter 2.3 to generate a plasmid map. The students will also use a computer program to generate a restriction map using a test DNA sequence. C. Terms Plasmid map

A representation of the relative distance of restriction enzyme cut sites of a plasmid.


An enzyme that targets and degrades RNA.

D. Background 1. Plasmid Maps Plasmid maps are easy-to-use tools to understand the features and components of a plasmid (see Figure 2.4.1). Construction of an accurate map of the sites where restriction nucleases cleave DNA is critical for almost all subsequent manipulations of genetic material. Restriction maps can be generated by restriction mapping (in case the DNA sequence s unknown) or via a suitable computer program (in case the DNA sequence is known).

31 2. Generating a Plasmid Map A. Computer Prediction If the sequence of a stretch of DNA is known, then it is possible to identify the restriction enzyme sites present in that piece of DNA using a computer program. There are many such programs available: some are free to use and some are not. In this class, we will use the tool hosted at the University of Colorado‟s website. A plasmid map generated this way can be used in many applications. For example, it can be used to determine whether a particular enzyme is suitable for a cloning project, or to generate a prediction to match Figure 2.4.1: Plasmid map deduced against actual experimental results. Since the after restriction digestion. selection of the appropriate cloning site is critical to the success of a cloning project, the ability to visualize the distribution of restriction sites and to be able to choose the correct ones to use is very important in gene cloning processes. B. Restriction Digestion If the entire DNA sequence is not known, a restriction map can be generated by performing restriction digestions and examining the lengths of the products produced under different conditions (i.e. by using restriction endonucleases with known sequence preferences). This is typically done by a combination of single (one restriction enzyme) and double (two restriction enzymes) digestions. By examining the pattern of digested products that we obtained by using two different single digestions (in our case EcoRI or HindIII) and a double digestion (EcoRI and HindIII simultaneously), we can piece together the original form of the plasmid using logical deductions. 3. RNAse RNAse is an enzyme that targets RNA molecules and destroys them. In this experiment, we are only interested in the plasmid DNA. Due to similarities in chemical properties of RNA and DNA, our procedure in Chapter 2.3 will have pulled down a substantial amount of RNA as well as plasmid DNA. To reduce the degree of RNA contamination in our analyses of restriction digestion patterns of the plasmid, we add RNAse to digest away the unwanted RNA.

32 E. Procedure

Each Student

Part I: Computer Prediction 1. Log on to D2l and look under the folder 2.4 for a MS Word file called “Unknown DNA.” Download this file and open it in MS Word. 2. Go to r/index.html 3. Scroll down and you should see two windows like the ones shown in Figure 2.42. 4. Select the entire sequence from the MS Word file, copy it, and paste into the top box. Figure 2.4.2. Using “Mapper” Program 5. Click “Create Map,” and the result should show up in the black window below. 6. Use the drop-down menu under the “Create Map” to change from “All restriction enzymes” to “Core Set of Enzymes.” 7. The short, vertical lines represent cut sites for a specific enzyme. The horizontal w h i t e l i n e Figure 2.4.3. Results of Mapper program on a demo piece of DNA represents your DNA sample. Clicking at different parts of the white line will give you different information concerning the fragment you just clicked on. In the example shown in Figure 2.4.3, the white line on EcoRV between the two vertical lines has been clicked, and the result displayed that “Clicked within a 76 bp EcoRV fragment on base 322.” This means that the length of the white line between the two vertical lines is 76 basepairs, and that the point where the mouse clicked on is base pair #322. 8. If you click on the name of the enzyme, you will see the recognition sequence, as well as the type of cut (5‟ overhang, 3‟ overhang, or blunt).

33 Part II: Restriction Digestion of plasmid (which plasmid do I have?). Each pair of students 1. Between you and your partner, pick one sample to use („A‟ or „B‟). Save the other sample. 2. Set up the following digestions: Table 2.4.1. Mixing chart for restriction digestion of DNA.

Component DNA (~1 µg) EcoRI HindIII RNAseA Buffer (10x) Sterile Water Final Volume

#1 µl 0.5 µl 0 µl 0.5 µl

#2 µl 0 µl 0.5µl 0.5 µl

#3 µl 0.5 µl 0.5 µl 0.5 µl

#4 µl 0 µl 0 µl 0 µl

25 µl

25 µl

25 µl


3. Incubate your reactions at 37 oC for 30 minutes. Next class period: 4. Prepare an agarose gel, following the instructions from Chapter 2.2. Use the 8 well combs. 5. Create 4 spots, each of 10 µl, and add 2 µl of loading dye. 6. Load samples onto the wells of the gel. Per table only load one of the undigested controls! 7. On each gel, include a 2-log DNA ladder. 8. Close the lid and make sure that the wells are closer to the cathode (negative charged electrode, -ve). 9. Set the power supply current level to low, the display to mV, and set the voltage to 200 mV. Check to see that bubbles are coming out from the two ends – this will tell you that the unit is properly connected. 10. Run the gel for 20 minutes. 11. Make a picture of your gel.


Chapter 2.5 Ligation of DNA A. Introduction Cutting (restriction digestion) and joining (ligation) of DNA molecules allows us to create new DNA sequences (e.g. plasmids) from different pieces of DNA, provided that they have compatible endings. The enzyme ligase is essential for joining DNA molecules in vitro. When termini that were digested by restriction endonucleases that create cohesive ends associate, the ends have nicks a few bases apart on the opposite strands. DNAse ligase can then repair these nicks to form an intact duplex. Ligase, the enzyme that enables this reaction, is in vivo involved in DNA replication and DNA repair. B. Objectives In this exercise, students will perform a ligation reaction using the digested products from Chapter 2.1. The product generated in this Chapter will be used in Chapter 2.6 for transformation. C. Terms Ligation

The process of joining two pieces of DNA together. Mediated by ligase.


Discontinuity in a double-stranded DNA molecule where there is no phosphodiester bond between adjacent nucleotides of one strand, typically through damage or enzyme action.

Cohesive End

A single-stranded end of a linear duplex DNA molecule which can hydrogen-bond with a complementary single-strand base sequence from the end of the same or another DNA molecule

Blunt End

A fragment of DNA resulting from the breaking of DNA molecule in which there are no unpaired bases or overhangs in the end.

D. Background 1. Ligation The success of a ligation reaction depends on several factors. First, the pieces of DNA to be joined should have cohesive compatible ends or blunt ends. Second, there should be enough material in a reaction to facilitate the random occurrence of having the ends react with each other, because while ligase can seal up pieces of DNA, it does not bring the pieces together. Rather, the meeting of end parts on a piece of DNA with the end parts of another piece of DNA is a function of thermodynamics and chance. Finally, the purity of the beginning products will affect the ligation reaction, as well.

35 E. Procedure

Each Pair

1. Retrieve your digested plasmid sample that had been frozen (product of Chapter 2.1, pKG15.3!). 2. Place the thawed tube of digested plasmid in the 75 oC heating block for 20 minutes. At the end, briefly centrifuge the tube to collect any condensations in the tube. 3. Set up the ligation reactions as follow: Table 2.5. Mixing chart for ligation.

Component Digested plasmid 10X Ligase buffer DNA Ligase Water Total

#1 (Control 1) 5 µl

#2 (Control 2) 0 µl

#3 (Ligation) 5 µl

0 µl

1 µl

1 µl

20 µl

20 µl

20 µl

4. The reaction will take place for 24 hours at room temperature, and then it will be stored for use in Chapter 2.6.


Chapter 2.6 Transformation of E. coli A. Introduction Once a new plasmid is created, we can shuttle it to a new bacterial host using transformation. Most bacteria have a natural tendency, called competency, to take up foreign DNA. Cells can also be prepared in specific ways to enhance the natural competency so that transformation can be more efficient. Selection of bacteria (typically by using an antibiotic for which resistance to its toxic effect is encoded on the plasmid) follows transformation to allow us to pick out the bacteria that have successfully received our plasmid. The most common selectable markers are ones that encode resistance to antibiotics such as ampicillin (ApR), chloramphenicol (CmR), tetracycline (TcR), and kanamycin (KmR). Other selection strategies include positive selection by inactivation (e.g. the ccdB gene which is lethal to host cells) and visual examination of reporter genes such as the lacZ gene product. LacZ encodes for -galactosidase which is easily detected by the occurrence of blue color if cells are grown on a medium that contains the chromogenic substrate Xgal (5-bromo-4-chloro-3-indolyl-beta-Dgalactopyranoside). In all transformation experiments, the identity of the plasmid needs to be confirmed after resistant colonies have been identified on selective growth medium. This is an essential step because not all ligated plasmids may be what we want, and other cloning artifacts (e.g. linearized plasmid integrating into the host chromosome) can arise that leads to resistant colonies that do not contain our desired plasmid. B. Objectives In this exercise, students will transform the plasmids they have generated in Chapter 2.5 into bacteria, select for transformants (bacteria that have successfully taken up the plasmid), and confirm the identity of these plasmids. C. Terms Competency

The ability of bacteria to take foreign DNA into their cells.

D. Background 1. Transformation There are many protocols for transformation, each specific to the need and requirement for different experiments. Eukaryotic cells can also be made to take up foreign DNA, even though they do not have the natural tendency to do so, but that process is called transfection. Transformation refers to introducing foreign DNA into bacteria.

37 The two most commonly used transformation protocols are electroporation and heat shock of chemically competent cells. In electroporation, foreign DNA enters into bacteria as a high current of electrical energy is passed through the cells. In using chemically competent cells, bacteria are prepared in special steps to enhance their natural ability to take up foreign DNA. The two methods are comparable, but they do differ in the preparation work required as well as the transformation rate that results. In this lab we will be using chemically competent cells. The two key factors that can affect the efficiency of transformation in using chemically competent cells are (1) proper handling of cells and (2) proper heat-shock procedure. Competent cells are prepared and then stored at –70oC to stop their metabolic activities. It is important to keep them from reaching room temperature because once they warm up, these cells rapidly lose their competency. Therefore, while handling the cells, be sure not to hold it in your hands directly and avoid exposing it to room temperature. The second critical step is the heat-shock treatment. To make the cells take up the foreign DNA more readily, we perform a heat-shock on the material. This means a short burst of heat (42oC) is applied, and then removed. The sudden increase in temperature causes a phase transition in the plasma membrane of the bacteria (making them more “leaky”) which facilitates the uptake of plasmid DNA. The proper time and temperature of the heat shock is very important to successfully transform bacteria and improper heat-shock can severely reduce the transformation efficiency. 2. Selection The antibiotic to use in selection will depend on the marker carried by the plasmid. In this Chapter, the plasmid carries a gene that conveys resistance to ampicilin. We will plate out the transformants onto nutrient agar plates containing ampicilin, so that only bacteria that harbor our plasmids will grow. See Chapter 2.5 for more information on plasmids and selectable markers. 3. Confirmation After we retrieve the survivors from the selection plates, we will need to confirm whether they truly carry a plasmid and to identify the plasmid. We will follow procedure in Chapter 2.3 to isolate the plasmids, and then perform restriction digestion followed by agarose gel electrophoresis (see Chapter 2.4).

38 E. Procedure

Each Pair

Part I: Transformation 1. Retrieve your frozen ligation product. 2. Check on the competent cells for thorough thawing by gently, but briefly, flicking them with your fingers. Do NOT let the cells warm up to room temperature at any time. 3. While waiting for the cells to thaw and label 3 clean 1.5 ml Microcentrifuge tubes (one for each of your ligation reactions) and place them on ice to pre-chill. 4. Once the cells are thawed, distribute 25 µl of cells to each of the 3 ligation tubes. 5. Add 2.0 µl of your ligation reaction, separately, to each tube (see page 35). 6. Keep the tubes chilling on ice for 10 minutes. 7. Bring the ice bucket with your samples over to the 42 oC water bath, and heat shock your samples for 90 seconds. 8. Return the samples to ice and let sit for 5 minutes. 9. Add 1 ml of sterile LB broth to each of the tubes, mix, and then set in 37oC shaking incubator. 10. Incubate for a minimum of 20 min. 11. While waiting for your incubation to end, label LB/Amp plates. You should label 2 plates for ligation condition #3, then one plate for ligation #1 and #2 each, for a total of 4 plates. 12. Add 150 µl of the appropriate ligation mixture per plate, and use the sterilized glass rod to spread the mixture until it is evenly distributed on the plate. 13. The plates will be incubated at 37 oC for 24 hours and then stored at 4oC for you. Part II: Inoculation 1. Choose 4 random colonies and inoculate them into LB broth. The culture will be grown overnight at 37oC with shaking. 2. To inoculate broth, use the provided sterile toothpick to touch the surface of the colony lightly. Even if you do not see any bacteria on your toothpick, as long as it made contact with the colony, there will be enough bacteria for the inoculation. Be sure not to allow the toothpick to touch anything other than the colony and the broth. 3. Wash the toothpick tip with the broth in the test tube provided. Each colony should be inoculated into a separate tube. Be sure to use a new sterilize toothpick between each colony. 4. Remember to label all your tubes.

39 Part III: Plasmid Isolation, Digestion, and Gel Electrophoresis 1. Using the cultures you inoculated the day before, isolate plasmids (from 4 different colonies) following protocol in Chapter 2.3. 2. Perform the following double digestions: Table 2.6. Restriction digestion of isolated plasmids.

Component/Colony DNA EcoRI HindIII RNAseA 10X Buffer Water Total

#1 10 µl 0.5 µl 0.5 µl 1 µl 2.5 µl 10.5 µl 25 µl

#2 10 µl 0.5 µl 0.5 µl 1 µl 2.5 µl 10.5 µl 25 µl

#3 10 µl 0.5 µl 0.5 µl 1 µl 2.5 µl 10.5 µl 25 µl

#4 10 µl 0.5 µl 0.5 µl 1 µl 2.5 µl 10.5 µl 25 µl

3. Incubate your digestions at 37 oC for 30 min. Next class period: 4. Use the gel you will have at your table (8-well comb). Receive your samples. Pick 3 out of 4, add 2 µl of the 5 x loading dye to each of the three digestion reactions, and load the entire sample onto the agarose gel. 5. On each gel, include 1 lane of the 2-log DNA ladder 6. Close the lid; make sure that the wells are closer to the cathode (negative charged electrode, -ve). 7. Set the power supply current level to low, the display to mV, and set the voltage to 200 mV. Check to see that bubbles are coming out from the two ends – this will tell you that the unit is properly connected. 8. Run the gel for 15 – 20 minutes. 9. Prepare a picture of your gel.


Chapter 3 Polymerase Chain Reaction A. Introduction The Polymerase Chain Reaction (PCR) is a rapid procedure for in vitro enzymatic amplification of a specific segment of DNA. PCR has spawned a multitude of experiments that were previously impossible. Dr. Kary Mullis first conceptualized the idea of the polymerase chain Figure 3.1. Exponential accumulation of product reaction (PCR) in 1983 and the during amplification Nobel Foundation awarded Dr. Mullis the Nobel Prize in Chemistry in 1993. PCR is characterized by the use of a heatstable DNA polymerase in a repeated cycle of heating and cooling to achieve logarithmic accumulation of the desired product. The initial enzyme used was the Klenow fragment of E. coli DNA polymerase I, which had to be freshly added during each cycle because the enzyme was heat inactivated during the heating cycle. The introduction of thermostable Taq DNA polymerase from Thermus aquaticus facilitated automation of the procedure. There are three nucleic acid segments: the segment of double-stranded DNA (dsDNA) to be amplified, and two single-stranded oligonucleotide primers flanking it. Additionally there is a protein component (a DNA polymerase), appropriate deoxyribonucleoside triphosphates (dNTPs), a buffer, and salts. PCR has become ubiquitous in both research and testing laboratories. This procedure allows us to make a large number of copies of a defined region of DNA starting with a small quantity of material. Applications of this procedure have extended from testing whether a subject carries a specific gene to whether food samples are contaminated by bacteria. B. Objectives In this Chapter, students will perform PCR to detect the presence of the human a1AT gene. C. Terms Primer Pairs

Primers are short, single-stranded DNA sequence that anneal to a defined region of DNA.


Forming of base-pairs between a primer and target DNA.



Separation of DNA double strands into single strands.


Process of DNA polymerase synthesizing new material.


Structures formed by two primers base-pairing with each other instead of with the intended DNA target.


Structure formed when a single-stranded primer forms base pairing with itself to make a loop.


Primer Melting Temperature (Tm). The temperature at which one 50% of the DNA duplex will dissociate to become single stranded.

D. Background While setting up a PCR is a very straight-forward procedure, the reasoning and analyses that went into designing a successful PCR is more complicated. Many PCR reactions work in a robust manner, i.e., they tend to give you the right amplicon under many different conditions. However, some PCR reactions are pickier, and require finetuning and trouble-shooting to optimize. Many important variables can influence the outcome of PCR. For example careful titration of the MgCl2 concentration is critical. Additives that promote polymerase stability and process sensitivity and strategies that reduce nonspecific primer-template interactions generally improve amplification sensitivity. Here are some of the major factors that can affect the quality of your PCR. 1. Integrity of template Even though PCR applications can amplify your region of interest from a small amount of material, the efficiency of the reaction will be improved by better quality of starting DNA. Quality in this context refers to contamination by proteins and/or chemicals as well as whether the DNA is sheared and broken. The better the quality of your starting DNA, the easier it is to get good PCR results at the end. 2. Primer design Primer pairs define the region that is to be amplified. The extent of base-pairing between the primers and the target has significant impact on the quality of PCR. Events that reduce the primer‟s melting temperature, often denoted as Tm, will reduce the specificity of amplification. Depending on the experimental question, this may be a desired result, but in general, lowering the Tm will lead to more unspecific amplifications. Many computer programs are available to aid in the design of primers (e.g. Here are some general guidelines for primer design: a) primers should be at least 15 base pairs long, b) have at least a G/C content of 50%, and c) anneal at a temperature in the range of 50 ºC to 65 ºC. Factors that affect the Tm of a primer include:


Extend of basepairing

Both the length and the composition of a primer will affect the extent of base-pairing. A primer that is 19 bases long will have a higher Tm than one that is 12 base pairs, in general, and a primer that has a good mix of GC/AT pairing will work better than primers that are overly GC or overly AT.

Secondary structures

Since primers are single-stranded products, they have a tendency to form secondary structures. In some cases, these structures can persist during amplification and lead to reduced efficiency. Primerdimers and primer hairpins are the two most common types of secondary structures.

3. Cycling conditions Thermocyclers are machines that can heat and cool the PCR samples in a rapid manner. The newer generations of machines are equipped with a heated lid, so that the samples will not form condensates on the lid during the repeated heating and cooling. A typical PCR cycling protocol consists of 4 main stages. Table 3.1: PCR cycling conditions Stage Temp. Initial Melting 96oC

Melting Annealing

96oC varies


72 oC

Final Extension Hold

72 oC 4oC

Duration Function 10 min (1) To ensure that the template strands are thoroughly separated at the beginning; (2) To activate the DNA polymerases 30 s To separate strands in new products 20 s To allow primers to attach to the correct sites. The actual temperature used will depend on the specific primer pairs, but is usually between 45oC and 65oC. varies Different polymerases work at different speed, but in general, it takes 1 minute to create 1 kb of product. varies This is usually 3x the regular extension time to allow any incomplete products to finish. ~ Hold at 4oC indefinitely until samples are collected. The low temperature prevents further polymerization.

The melting, annealing, and extension steps form the cycling stage, and they are repeated for 25 to 40 times.

43 4. Reagents In addition to the DNA template and the primer pairs, a PCR reaction requires the following components: DNA Polymerase

The polymerase used must be heat-stable so that it does not fall part upon repeated rounds of heating and cooling. There are two main types of polymerases: the proofreading type and the nonproofreading type. The proofreading type generates blunt-end products, and gives more accurate amplification. The other kind generates products with a dangling A residue and is usually cheaper.


Each polymerase comes from the supplier with its own reaction buffer, which contains various salts in a pH-controlled solution to optimize the performance of the polymerase.


A collection of the 4 different deoxyribonucleotides, (G,C,A,T) is need to provide building blocks for new DNA. They are usually in a pre-made mixture where each base is in equal proportion to the other three.

E. Procedure

Each Pair

Part I: Setting up PCR 1. Obtain the digested DNA and the undigested DNA from frozen storage (page 21, Table 2.1; pPGK15.3). 2. Obtain the PCR tubes (0.25 ml thin-walled) and label them. Due to the limited space on the tube to label, you will be assigned an alphabet to represent your group. 3. We are using a PCR pre-mix that contains all the components except for the template and the primer. 4. Set up the reaction as in Table 3.2.

44 Table 3.2: PCR mixing chart

Component Buffer mix (2x) Template Primer 1 Primer 2 Water Total




1 µl Undigested 1 µl 1 µl

1 µl Digested 1 µl 1 µl


25 µl

25 µl

25 µl

1 µl 1 µl

5. We will run your samples on the PCR machine and then freeze it for you until we run it on a gel. Part II: Electrophoresis 1. 2. 3. 4.

Prepare an agarose gel using the comb with 8 teeth. Add 2.0 µl of the 6x loading dye to 10 µl of each of your PCR samples. Mix well. Load 10 - 12 µl of each tube onto an agarose gel. Run the gel at about 200 mV for 30 min.

dNTPs Primer 1 DNA Polymerase DNA Primer 2

Figure 3.2. Components in the PCR reaction tube.



Chapter 4 Enzymatic Activity of Gene Products (Proteins) A. Introduction In this chapter you will be assaying the proteolytic activity of the two enzymes pepsin and trypsin. Both enzymes are peptidases that aid in the digestion of proteins by hydrolyzing the polypeptide chain into shorter peptides. Pepsin is found in the stomach of vertebrates whereas trypsin is secreted into the duodenum. Due to the very different conditions in the stomach and the duodenum both enzymes show maximal activity at very different proton concentrations (pH optimum). In order to follow the activity of both peptidases we will be using azocasein. Azocasein is a protein with a molecular weight of Mw = 23.6 dalton and consist of casein conjugated to an azo-dye. The azo-dye in the protein absorbs light at 440 nm (orange/red color). Azocasein serves as a general substrate for proteolytic enzymes. The assay is based on the fact that undigested azocasein can be denatured with TCA (trichloroacetic acid) and removed by centrifugation, whereas the digested peptides will remain in solution. The amount of red color in the supernatant after digestion and acid denaturation is therefore directly proportional to the proteolytic activity of the investigated enzyme.

B. Objectives In this Chapter the student will determine the effect of temperature and proton concentration on the activity of the two enzymes pepsin and trypsin. The rate of reaction will be determined by following the increase of red color. The student will be familiarized with the concept of how abiotic factors influence the reaction rates of cellular systems. C. Terms TCA

(Trichloroacetic acid): An analogue of acetic acid in which the three hydrogen atoms of the methyl group have all been replaced by chlorine atoms. TCA is a strong acid. In this chapter it is used to denature proteins and to halt all enzymatic processes.


A protein that serves as the substrate for the enzymes pepsin and Trypsin. Azocasein is conjugated with a red dye.


Pepsin is an enzyme in the stomach that degrades food proteins into peptides.


A serine protease (enzyme) found in the digestive system of many vertebrates that is secreted into the duodenum. Sodium hydroxide is a caustic metallic base used for alkalization of our sample.



D. Background Enzymatic activity is influenced by several abiotic factors such as temperature, ion composition, and pH value. For example, pH can have an effect of the state of ionization of acidic or basic amino acids. The state of ionization of amino acids in a protein can be altered by changing the proton concentration in the medium. This then alters the ionic bonds that help to determine the 3-D shape of the protein, leading to altered protein recognition or enzyme inactivity. In general enzymes have a pH optimum. However the optimum is not the same for each enzyme. Increases in the temperature of a system results from increases in the kinetic energy of the system. This has several effects on the rate of reactions: a) more energetic collisions; b) the number of collisions per unit time will increase; c) the heat of the molecules in the system will increase. As the temperature of the system is increased, the internal energy of the molecules in the system will increase. The internal energy of the molecules may include the translational energy, vibrational energy, and rotational energy of the molecules. Furthermore it includes the energy involved in chemical bonding of the molecules as well as the energy involved in nonbonding interactions. Chemists have a rule of thumb that a 10 °C increase in temperature gives a doubling of the reaction rate. This rule is loosely derived from the Arrhenius equation. This is defined by the Q10-value: Q10 = Ratio of reaction rates for a 10 °C change = Rate at 30 °C / Rate at 20 °C = ~2. Basically, as the temperature increases so does the kinetic energy of the reactants. An increase in temperature will therefore increase the rate of an enzymatic reaction. If this chemical potential energy increase is great enough some of the weak bonds that determine the three dimensional shape of the active proteins may be broken. This could lead to a thermal denaturation of the protein and thus inactivate the protein. Thus too much heat can cause the rate of an enzyme catalyzed reaction to decrease because the enzyme or substrate becomes denatured and inactive.

The Arrhenius equation:

k=A*e(-Ea/R*T) Where k is the rate coefficient, A is a constant, Ea is the activation energy, R is the universal gas constant, and T is the temperature (in Kelvin). R has the value of: 8.314 x 10-3 kJ mol-1K-1.

47 E. Procedure

Each Pair

You will be working in groups of 2, but your data will be pooled as a class. Please refer to the instructions in class to determine which assay condition is assigned to your group. Groups 1 to 3, will be assaying the activity of trypsin. Groups 4 to 6, will be assaying the activity of pepsin. For each group, there will be an A and a B team. Team A will be assaying pH 1, 3, 5, 7, and 9. Team B will be assaying pH 2, 4, 6, 8, and 10. You will be creating buffer solutions with different pH values according to Table 2. Buffers with pH 1, 9, and 10 are handed out by the TA. Part I: pH optimum of Trypsin and Pepsin (1st Lab Period)

Table 4.1. Assay conditions investigated by different groups. Gp # 1 2 3 4 5 6

Team A B A B A B A B A B A B

Enzyme Trypsin Trypsin Trypsin Trypsin Trypsin Trypsin Pepsin Pepsin Pepsin Pepsin Pepsin Pepsin

pH 1,3,5,7,9 2,4,6,8,10 1,3,5,7,9 2,4,6,8,10 1,3,5,7,9 2,4,6,8,10 1,3,5,7,9 2,4,6,8,10 1,3,5,7,9 2,4,6,8,10 1,3,5,7,9 2,4,6,8,10

1. Determine which condition your team of two people will be investigating. Using the table below (Table 4.1), create buffer solutions of the correct pH. 2. Confirm with the supplied pH-paper that your solutions have the correct pH. 3. From the pH buffer stock you just made, or the buffers that were handed out by the TA, aliquot out 250 µl into a fresh tube. Label the tube appropriately. 4. Add 250 µl of azo-casein to each tube (Note: This is your substrate your enzyme will act on). Keep mixtures on ice. 5. Retrieve the appropriate enzyme (trypsin OR pepsin) from the TA. Mix evenly. This starts your reaction. Time is running now. 6. Add 15 µl of enzyme to each tube. Keep

all reagents on ice. 7. Incubate your reactions at 37 ˚C for exactly 30 min. 8. Retrieve your samples, and add 350 µl TCA solution (Note: The addition of TCA will stop the reaction). 9. Centrifuge for 10 min at maximum speed to remove undigested casein. 10. Remove 700 µl of the supernatant containing colored casein fragments from each tube to a fresh tube. Discard the tube with the debris. 11. To each tube, add 230 µl NaOH solution. Mix well (Note: the addition of NaOH will increase the solubility of casein fragments and assure a uniform pH among the different samples) 12. Transfer the mixture from each tube to a disposable cuvette. 12. Obtain an OD reading at 440 nm. 13. Record the data (Table 4.2).

48 Table 4.2. Results (Part 1) and Mixing Chart. pH

Na2HPO4 (µl)

Citric Acid (µl)































OD440 nm

Part II: Temperature Optimum of Trypsin and Pepsin (2nd Lab period) Note: pH used for this experiment will be determined in Part I above You will be working in groups of 2, but your data will be pooled as a class. Please refer to the instruction in class to determine which temperatures are assigned to your group. You will assay the activity of both enzymes. Groups 1 to 3 will be assaying at temperatures 0, 15, 35. Groups 4 to 6 will be assaying the activities at 55, 75, and 95 ˚C. Gp # 1 2 3 4 5 6

Team A B A B A B A B A B A B

Enzyme Trypsin Pepsin Trypsin Pepsin Trypsin Pepsin Trypsin Pepsin Trypsin Pepsin Trypsin Pepsin

Temp ˚C 0,15,35 0,15,35 0,15,35 0,15,35 0,15,35 0,15,35 55,75,95 55,75,95 55,75,95 55,75,95 55,75,95 55,75,95


1. Make a fresh buffer stock. Aliquot out 250 µl into a fresh tube. Use the buffers that were found to have the optimal pH for the respective enzyme (Part I). Label the tube appropriately. 2. Add 250 µl of azo-casein to each tube. Mix evenly and incubate the mixture for 5 minutes at the desired temperature BEFORE adding the enzyme. 3. Retrieve the appropriate enzyme (trypsin and pepsin) from the TA. Add 15 µl of enzyme (either pepsin or trypsin) to the respective tube and place the tube directly back into the water bath or heating block. This starts your reaction. Time is running now (30 min!). 4. Incubate your reactions at the desired temperatures for exactly 30 min. 5. Retrieve your samples, and add 350 µl TCA solution. Note: The addition of TCA will stop the reaction. 6. Centrifuge for 10 min at maximum speed to pellet undigested casein. 7. Remove 700 µl of the supernatant containing colored casein fragments from each tube to a fresh tube. Discard the tube with the debris. 8. To each tube, add 230 µl sodium hydroxide (NaOH) solution. Mix well. 9. Transfer the reaction mixture from each tube to a disposable cuvette. 10. Obtain an OD reading at 440 nm. 11. Record the data (Table 4.4). Table 4.4: Results (Part 2)

Temperature (˚C) 0 15 35 55 75 95

Pepsin OD440 nm

Trypsin OD440 nm


Chapter 5.1 Extra Credit Lab - Cell Counting Contest A. 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. Near the beginning of the last century R. G. Harrison was experimenting on frog nervous tissue. He believed nerve fibers grew from individual neurons. In 1907, he placed a piece of neural tube tissue from a frog embryo into a drop of clotted lymph on a cover slip. He observed that axons did indeed arise from individual neurons. His findings helped establish the concepts of nervous system structure that are still accepted today. He was the first to show that cells could Fig. 5.1.1: The cell cycle. 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 two 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 are a common 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 Chapter, 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 culture or 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

51 the culture flask). Non-adherent cells, 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 of culture can be taken simply mixing the flask. For the counting contest you will be supplied with two different animal cell lines. Sf-21 cells were derived from ovary cells of the fall armyworm Spodoptera frugiperda and Kc167 cells from embryos of the fruit fly Drosophila melanogaster. B. Objectives In this chapter, students will learn the use of hemocytometer. This lab prepares the student for the following sections in which the student will design an experiment that investigates the impact of several abiotic factors on growth and viability of Drosophila melanogaster Kc167cells. C. Terms Hematocytometer

Chamber to estimate the concentration of cells.

Trypan Blue

Cell impermeable dye used to determine membrane integrity (viability).

D. Background 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 5-2). 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 mm x 1 mm squares (Figure 5-3). 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 5-3)] is 1.0 mm x 1.0 mm x 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. Think of it like this: If you made a 10 by 10 array of these counting chambers (to get 100 of them) and stacked these 0.1 mm tall “pancakes” 100 high, you would have a total of 10,000 counting chambers and their combined volume would be exactly 1 cm3, or 1 ml.

52 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.

Fig. 5.1.2: The hematocytometer.

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 under filling 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 next. 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 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 mm 2 square contains between 25250 cells (i.e., a beginning concentration of 25 x 104 to 250 x 104 cells/ml). For statistical reasons, more than 200 cells should be counted (the counting error is approximated as the inverse of the square root of the total count). When cells are touching a line, it is

53 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.

Fig. 5.1.3: 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 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 cannot 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) x 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.

54 E. Procedure

Each Pair

You will be working in groups of 2, but your data will be assessed as a table. For each group, there will be an A and a B team. Team A will be counting sample „A‟ and the B team sample „B‟. You will count at least 200 cells of each sample. One sample will consist of Sf-21 cells (~20-25µm diameter) the other sample of Kc167 cells (~5-8µm diameter). You will not know which team got the Sf-21 cells and which team got the Kc167 cells. You have to judge the cell line by comparing the sizes. Your cell counts will be collected on the white board. 1. Obtain samples „A‟ and „B‟ 2. Mix your sample by brief vortexing. 3. Load onto a small piece of parafilm and mix: 15 µl of the cell suspension and 15 µl 0.4% (w/v) trypan blue solution and wait 30 seconds. 4. Add 20 µl 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 10-3) to fit into the field of view 5. From this point there are two ways to count cells and calculate cell densities using a hemocytometer: 6. Large cells: If the number of cells visible in the large central square (square “A”) is less than about 100 (make a ball-park estimate), count the number of cells in the entire large central 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 all the cells in the square have been counted. When finished counting, you will have counts for 5 large squares. Switch to the other side of the hemacytometer (to the other counting chamber) and repeat the process, again counting the number of cells in each of 5 large squares. Recall that with the cover slip in place, the volume of one large square is 1 x 10-4 ml (1/10,000 ml). Cell density is calculated as: The # of cells counted / number of (big) squares counted (1 square/10 -4 ml) x dilution/concentration 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) x 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 to accurately reflect the cell density in the original culture. On the other hand if you take 1 ml of cell suspension, pellet the cells by centrifugation, then bring them up in a total volume of 100 μl, 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

55 be multiplied by 0.1 (i.e., divided by 10) to correct for the act of concentrating the cells. 7. Small Cells: If a very large number of cells are visible (over 200 cells in grid “A”), or if the cells are very small count only the cells in one small grid of corner grids. In practice, count the cells in each of the 16 small squares located within the larger corner squares. 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 10 cells, but do not stop counting until the count of a given (small) square is complete. Count all 16 small squares in a corner grid. Using the stage adjuster, move to the other counting chamber and count the number of cells the opposite corner grid. Cell density is calculated as: The number of cells counted x 16 (1 square/10-4 ml) x dilution/concentration factor = number of cells / ml in the original culture. Cell viability is calculated as: The number of living cells (number of dead cells + number of living cells) x 100 = percent viability. Extra Credit 1: Which sample contained Sf-21 cells and which contained Kc167 cells? (1 pts) Extra Credit 2: What are the life and dead counts for sample A and B? Each student of the group that is nearest to the correct number will get 2 pts extra credit.


Chapter 5.2 Sterile Cell Culture: Experimental Designing A. Introduction Last lab you learned how to determine the number of cells in a cell sample. This week you will design an experiment that investigates the impact of several factors on the growth of the Drosophila melanogaster cell line Kc167. You will have two different Kc167 cell lines for your experiment available. The parental wild-type Kc167 cell line, and Kc167 cells that were transfected with a GFP tagged Late Embryogenesis Abundant (LEA) protein from the brine shrimp Artemia franciscana (Kc167-LEA1-GFP). Since you are now going to culture your cells it is important to realize that you have to perform your cell culture experiment in an environment that is as sterile as possible. Some groups will choose to perform the experiment in a vertical flow hood. Other groups will perform the experiment on the bench and make sure that cell culture dishes are only open for brief amounts of time and that the working surface was sterilized with 70% ethanol. In a vertical hood for cell culturing air is filtered 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 towards the operator in a horizontal flow across the work surface. 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. Thought they look similar, fume hoods and sterile hoods have fundamentally different functions. Fume 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. Starting from the rear of the hood and working forward, swab down the work surface liberally with 70% ethanol. 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, and then return it to the hood, so that the next users think they are sterile. Never do this. Leave the tips in the hood.

57 4. Securely cap all containers before removing them from the hood. Containers should remain securely capped at all times when outside 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. Do not work close to the front of the hood where sterility may be compromised. 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. In this chapter, you will culture and propagate Kc167-LEA1-GFP cells and assess their growth rate under various conditions. The growth rate of the transformed cells will be compared to that of the wild-type Kc167 cells. B. Objectives In this chapter, students will be introduced into sterile cell culture and design an experiment that investigates the impact of several factors (temperature, osmotic stress, salt stress, light) on the growth of wild-type and transfected Kc167 cells. C. Terms FBS

Fetal Bovine Serum. A serum prepared from the blood of a fetal calf. Widely used in cell culture media.

12-well dish

Cell culture vessel.

LEA protein

Late Embryogenesis Abundant protein. Class of proteins that is expressed in desiccation tolerant organisms.


Widely used cell culture medium for insect cells.


Cells harboring a gene that has been introduced using genetic engineering.


Green Fluorescence Protein. Protein composed of 238 amino acid residues (26.9kDa) that exhibits bright green fluorescence when exposed to blue light.

58 D. Background The vast majority of animal species do not tolerate severe water stress, but the encysted embryo of the brine shrimp Artemia franciscana is an exceptionally useful organism to investigate physiological mechanisms for enduring extreme environmental insults. Any substantial reduction in cellular water poses a threat to survival. Nevertheless anhydrobiotic animals survive virtually complete loss of cellular water. The mechanisms that govern “life without water” (anhydrobiosis) are still not well understood. With certain exceptions, it seems that a recurring strategy for tolerating severe water loss involves the accumulation of highly hydrophilic macromolecules such as late embryogenesis abundant (LEA) proteins. Insect cells are useful to investigate possible function of LEA proteins. You will be designing your experiment to investigate the impact of different growth conditions on viability and proliferation of Kc167 and Kc167-LEA1-GFP cells. E. Procedure

Each Table

A. Design your experiment as a group after meeting with your advisor. Bring your experimental design with to your group meeting. B. Factors that you may investigate: 1. Concentration of potassium chloride (KCl): 0 mM up to 1000 mM in 200 mM increments. 2. Concentration of sodium chloride (NaCl): 0 mM up to 1000 mM in 200 mM increments. 3. Concentration of sucrose: 0 mM up to 500 mM in 100 mM increments. 4. Temperature (4 ˚C, 20 ˚C, 37 ˚C). 5. A combination of 1-4. C. You will start your experiment in the first lab section. You will measure cell growth and viability by Trypan Blue exclusion. In order to gain a solid amount of data you will be required to perform cell counts outside the class period on day 2 and 5. You will also be able to assess the expression level of GFP by fluorescence microscopy. More detailed instruction will be given during the course. Use the figures on the next page to plan your plate layout. Turn in your experimental layout during the group discussion (Assignment 2, 15 points)



Chapter 6 Western Blotting A. Introduction The term Western blotting refers to procedure which does not directly involve nucleic acid, but which is of importance in gene manipulation and proteomics. Western Blot is a method for detecting a specific protein in a mixed protein sample. It shares with Southern blots the core idea of separation of components in a sample by size, followed by transfer onto a membrane, and then detection with a probe specific to the target. The first step is to transfer or blot the pattern of separated proteins from a polyacrylamide gel onto a sheet of nitrocellulose. An efficient method of transfer is achieved by electroblotting. A sandwich of gel and nitrocellulose is compressed in a Figure 6.1. Protein detection principle of cassette and immersed, in a buffer, between two Western blotting using horse radish parallel electrodes. A current is passed at right peroxidase (HRP). angles to the gel and separates the proteins out of the gel and onto the nitrocellulose sheet. The nitrocellulose membrane with bound proteins is referred to as blot. The blot is than incubated in a dilution of antiserum (1º Ab, primary antibody) directed against the protein of interest. By using a secondary antibody (2º Ab) that is appropriately labeled (e.g. horse radish peroxidase, HRP) and binds to the primary antibody the interaction can be visualized. B. Objectives In this chapter, students will isolate protein from different types of fish muscle, separate the total protein on an SDS-PAGE and then detect the presence of myosin. C. Terms Sodium Dodecyl Sulfate (SDS)

An organic, anionic detergent used in denaturing proteins.

Polyacrylamide Gel Electrophoresis (PAGE)

Separation of protein samples using a thin polyacrylamide gel. Polyacrylamide is a potential neurotoxin, so the material should be handled while care while wearing gloves.

Primary Antibody

Antibody against the target protein to be detected.

61 Secondary Antibody

Antibody raised against the part of the primary antibody that is unique to the primary host species.

Coomassie Blue

A staining solution that will stain all proteins in a sample.


Tris-buffered saline with Triton X100. TBS is a saline solution for washing membranes, and Triton X100 is a non-ionic detergent used to prevent antibodies from binding non-specifically to the membrane.

Thiol Reagent

Dithiothreitol (DTT) or mercapotoethanol are commonly used to reduce disulfide brides between cysteins in proteins.


Substance that causes the immune system to produce antibodies antibodies against it.

D. Background Unlike Southern and Northern Blots, which rely on hybridization of nucleic acids, Western Blot relies on protein-protein interactions for detection. The quality of the primary antibody is significantly more difficult to control than the quality of DNA/RNA probes is. 1. Protein Separation Proteins can be separated under native or denaturing conditions. Under native conditions, proteins will separate out based on both overall charge distribution on the protein surface and three-dimensional shape. Under denaturing conditions, such as SDS-PAGE, proteins are separated based on their linear length. The SDS acts to neutralize the uneven charges on the protein by creating a relatively linear charge-tolength ratio and also by unfolding the protein. When denatured by heating in the presence of excess SDS and a thiol reagent, most polypeptides bind SDS in a constant weight ratio such they have essential identical charge densities and migrate in polyacrylamide gels of the correct porosity according to the size of the polypeptide. 2. Transfer The transfer of proteins from a polyacrylamide gel onto a membrane requires more energy than the capillary transfer method employed in both Southern Blots. Instead of relying on the capillary action of solvent, an electric current is applied to mobilize the proteins onto the membrane. 3. Antibodies When animal systems are challenged by a foreign substance, part of the response is to generate immunoglobulin G (IgG) that can target these foreign substances. By nature,

62 these IgG tend to be rather specific to the foreign objects (antigen) that they target. Capitalizing on this response, researchers can inject the target protein to be detected into an animal host to elicit IgG production. These IgG can then be harvested and used as primary antibodies. There are some considerations for this process: a. The primary host animal should not be the same species as the target protein is from. That is, if we want to detect the gene for breast cancer in rats, we should not use rats as the primary host. The most common primary hosts animals include mice, rats, rabbits, and chickens. b. Typically, only a portion of the target protein is used in the injection. This enables the researchers to choose only the region on the target protein that separates it from all other proteins so that the antibodies will not cross-react with non-targets. The selection of which portion of the protein to use can be tricky, and it depends on the uniqueness of that region as well as the likelihood of that region being on the surface of the protein instead of being buried during 3-D folding. c. Primary antibodies can be either polyclonal or monoclonal. When antibodies are first generated, they are polyclonal, meaning that there are probably multiple versions of IgG that react to the same foreign protein, with each version interacting with a different region of the target. It will also contain IgG that do not interact with the target at all, but which are simply co-purified. Researchers can take the polyclonal antibodies and generate a pure form of the IgG, where only one variant is present. Monoclonal antibodies are made by identical immune cells that are all clones of a unique parent cell. These antibodies are specific to the target protein and pure in composition. Obviously, the increase in specificity of monoclonal antibodies comes with an increased price tag. Secondary antibodies are generated by injecting the region of primary IgG that is specific to the primary host species into a secondary animal host. The secondary host needs to be of a different species than the primary host, e.g. if the primary antibody host is a rabbit, then the secondary antibody host cannot be a rabbit. The secondary antibody is purified and then conjugated to a reporter enzyme, similar to how Southern probes are linked to reporter enzymes. E. Procedure Part I: Protein Isolation and Separation 1. 2. 3. 4. 5. 6. 7. 8. 9.

Obtain fish muscle sample. Put the fish muscle sample into a 1.5 ml Microcentrifuge tube. Add 250 µl of Laemmli sample buffer to your tube. Close lid and flick the tube 15 times. Incubate at room temperature for 5 minutes. Transfer the liquid portion to a new, clean tube. Heat for 5 minutes at 95oC to denature protein. Let sample cool off to room temperature right before loading. Load 10 µl of sample into the gel.

Each Pair

63 10. The SDS-PAGE will run for 1hr at 200V. Part II: Western Blot Washes 1. Prior to this period, the protein will have been transferred onto a membrane for you and put through the blocking step. 2. Obtain your sample in a 10 ml round-bottom snap-cap tube. 3. Briefly rinse the membrane with 5 ml of 1X TBS-T buffer. 4. Add 5 ml of mouse anti-myosin antibody (1o Ab) and place on rolling incubator for 15 min. 5. Briefly rinse the membrane twice in 5 ml TBS-T. 6. Wash membranes twice in 5 ml TBS-T, for 5 min each. 7. Decant the solution. 8. Add 5 ml of goat-anti-mouse antibody (2o Ab) and place on rolling incubator for 15 min. 9. Briefly rinse the membrane twice in 5 ml TBS-T. 10. Wash membranes twice in 5 ml TBS-T, for 5 min each. 11. Decant the solution. 12. Add 5 ml of developing agent (horseradish peroxidase detection agent) and allow the color to develop fully. This usually takes 15 to 30 minutes. 13. Rinse the membrane twice in water. 14. Remove membrane and pat dry. 15. Hand your membrane to the TA to be photographed.


Chapter 7 Gene Reporters A. Introduction In nature, most organisms do not express every single gene they possess at all time. The regulation of when to “turn on” a gene, for how long, to produce how much proteins, is critical to the success of living organisms to adapt to a changing external environment. In multi-cellular eukaryotes that are composed of different tissue types, regulation of gene expression is required for tissue differentiation, e.g., genes specific to the function of neurons are not expressed in liver cells, or vice versa. One of the tools used in studying the regulation of gene expression is called a reporter gene system. There are two main uses for reporters: (1) to reveal the localization characteristics of a protein in the cell and (2) to reveal the activity of the gene promoter. B. Objectives In this chapter, students will perform both qualitative and quantitative assays for the reporter -galactosidase controlled by the human 1AT promoter to determine whether the hormone dexamethasone impacts promoter activity. C. Terms -galactosidase

An enzyme that cleaves  sugar bonds. E. coli for example produces the enzyme in the presence of lactose (β-Dgalactopyranosyl-(1→4)-D-glucose) to hydrolyze the disaccharide into the monosaccharide glucose and galactose.

5-Bromo-4-ChloroColorless chemical compound that creates a blue precipitate 3-Indolyl--D-galactoside when cleaved by -galactosidase. (X-Gal) Ortho-nitrophenyl--Dgalactoside (ONPG)

Colorless chemical compound that creates a fluorescent compound that will emit light at 405nm upon UV excitation.


A synthetic hormone known to stimulate expression of some genes specific to liver cells. It belongs to the glucocorticoid class of steroid drugs with strong anti-inflammatory properties and is several-fold more potent than the naturally occurring hormone hydrocortisone.


Phosphate buffered saline. A NaCl based solution with optimized pH and osmolarity used in mammalian cell culture.

65 BSA

Bovine serum albumin. Serum protein from cows with has numerous biochemical applications.


Bicinchoninic acid. Chemical that is commonly used to determine the concentration of protein in a solution.

D. Background For the qualitative assay, cells will be “fixed” (fixation of cells attached to the culture dish with organic solvents) to preserve -galactosidase activity. This step stops the overall metabolic activity of the cells, but does not inhibit the enzyme -galactosidase. Therefore, presence or absence of the enzyme can be demonstrated by incubation with X-gal solution. If the cells exhibit -galactosidase activity the development of color can be observed. However, this approach allows us only to demonstrate the absence or presence of the enzyme. We are not able to quantify the amount or level of activity that is present in our sample (and therefore we cannot quantify the activity of the 1AT promoter for which -galactosidase acts as reporter). In order to be able to estimate how much activity is present in the sample we need to apply a quantitative approach. For the quantitative assay, in order to compare our results more meaningfully, we have to perform a few standardization steps using control samples. A standardized unit of measurement for -galactosidase activity using ONPG was developed by Dr. Miller (1972). This unit was originally developed to be used with bacterial cultures: Miller Unit =

A420  1.75  A550 v  t  A600

A420: absorbance of enzyme assay product o-nitrophenol at 420 nm light A550: scattering of bacterial cell debris at 550 nm light  A600: absorbance of bacterial culture at 600 nm, used as an indicator of cell density v: volume in milliliters of the original cell culture used in the assay t: time in minutes of the assay Since we are not using whole bacterial cells, we can simplify the above formula to: A Activity = 405 qt A405 is the absorbance of enzyme assay product o-nitrophenol at 405 nm light q is the quantity of protein used in the assay, which is calculated as c*v  where c is the concentration of total protein (mg/ml) and V is the volume of sample in ml used in the assay t is the duration in minutes of the assay To determine c, we will need the help of a standard curve.

66 Protein Standard Curve The protein standard curve we are using is a graph we generate by plotting known quantities of protein (in µg/ml) against their respective absorbance at 562 nm after reaction with a detection reagent. (This assay is different from the enzymatic assay for ONPG. We will actually need the value of the protein assay to calculate the enzyme activity in the ONPG assay). In order to estimate the protein concentration in our samples we will be using a copper based protein detection assay. The BCA detection kit contains a reagent that will yield a color (purple) in the presence of protein. The more protein is present in the sample, the deeper the purple color. The BCA protein assay is a combination of the protein-induced biuret reaction with the colorimetric detection of the resulting Cu1+ cation by bicinchoninic acid (BCA). The reduction in light intensity of monochromatic light that passes through a solution is proportional to the intensity of the light I0 (incident light), the thickness of the sample dx, and a constant к (this constant becomes ε for a compound in solution). The absorbance is a useful parameter since it is proportional to the concentration of the absorbing species and the path length: A = -log (Im/I0)


A = ε • c • dx


Where ε is the molar extinction coefficient (M-1 cm-1), c the concentration of the absorbing species and dx the path length the light has to travel. In case the path length (in most cases 1 cm) and the molar extinction coefficient are known we can calculate the concentration of a compound based on Formula II. Absorbance is a dimensionless term between 0 and ∞. Due to light scattering and other secondary concentration depended processes that may occur at higher concentrations the measured absorbance should be between 0.01 and 1. However, we don‟t know the exact extinction coefficient of the reaction product between protein and BCA in the assay and therefore we have to perform a calibration curve using known quantities of protein. A relative cheap and commonly for this purpose used protein is bovine serum albumin (BSA). We will measure the intensity of the purple color at 562 nm at different BSA concentrations. By establishing a graph of known protein quantities and their respective level of color, we can estimate the quantity of unknown proteins by comparing their intensity against the standard curve.

E. Procedure

Each Group of 4

In this exercise, you will be working in groups of 4. From each group, 1 person will perform the Qualitative Assay Part 1, while the other 3 will work on the Quantitative Assay Part 1. In the next part, the group will work together to finish the two assays. For both types of assays, you will be working with 1 sample of FT (negative control without reporter-gene construct in the cell) cells incubated with 1x10-5 M DEX, and 3 samples of

67 FT-Beta cells. The 3 wells of FT02B cells will have been incubated with either 0, 1x10-5, or 1x10-6 M dexamethasone for several hours. I. Qualitative Assay Part: Fixation 1. Come up to the TA work bench. Your samples will be growing in a 24-well plate. Treat all your wells the same way. 2. Label on the provided sheet the wells you have chosen. 3. Remove the nutrient medium in the wells with a micropipette. You may have to tilt the plate slightly to get all the solution out. 4. Add 1 ml of fix solution (1% formaldehyde, 0.2% glutaraldehyde, in PBS). 5. Incubate at room temperature for 5 minutes. 6. Remove fix solution with a micropipette. 7. Rinse wells twice, each time with 1 ml PBS solution. Note: Add PBS gently to avoid dislodging the attached cells. 8. Remove PBS solution. 9. Add 0.5 ml of X-gal solution.

II. Quantitative Assay Part: Enzyme extraction 1. 2. 3. 4. 5. 6.

Decant the nutrient medium. Rinse plates twice, each time with 5 ml PBS Add 1 ml TEN buffer (40 mM Tris pH 7.4, 1 mM EDTA, 0.15 M NaCl). Incubate for 5 min. Dislodge cells by gently scraping the bottom of the plate with a scraper. Transfer everything from the plate into a 1.5 ml microcentrifuge tube using the 1000 l micropipette. 7. Microcentrifuge at 6000 rpm for 4 minutes to pellet cells. 8. Remove supernatant (liquid) with a micropipette. 9. Resuspend pellet in the same tube with 100 l 0.25 M Tris, pH 7.8. 10. Freeze sample. Next class period: 11. Thaw samples. 12. Centrifuge tubes at maximal speed for 5 min to remove cellular debris. 13. Use the samples in part III and IV.

68 III. Qualitative Assay Part: color development (fixed plate) Examine the degree of color development in each well. Record your observations: Table 7.1: Results (Qualitative).

Cell Type


Dexamethasone 10-5 M concentration





10-5 M

10-6 M


IV. Quantitative Assay Part: Generating a protein standard curve (thawed samples). 1. Pipette 10 l of each BSA sample into its own well in a 96-well microplate. The working range = 0, 100, 200, 400, 600, 800, 1000, 1500 μg/ml (Row A1-8). 2. Using fresh wells for each sample, prepare one well with 10 l and one well with 5 l of your protein extracts (Row B1 – 8). 3. Add 250 µl of Bradford reagent (e.g. Coomassie Plus) to all the wells. 4. Incubate at 30 min at room temperature. 5. Obtain a reading of the plate at 595 nm wavelength. Record your results. EXAMPLE CALCULATION FROM PREVIOUSE RESULTS: YOU NEED TO PLOT AND USE YOUR OWN STANDARD CURVE!


1.2 [BSA] vs OD 560nm Plot 1 Regr


OD 560



= 0.00XX*[Protein] + 0.0X

Curve 1: Coefficients: b = 0.02 m = 5.4e-3 r ² = 1

0.6 0.4 0.2 0.0 0






[BSA], g/ml

Fig. 7.2a. Example of a standard curve analysis to estimate protein concentration in an unknown sample.

Table 7.2b Results Protein Assay at ʎ= 595 nm.



[BSA] µg/ml

















Results OD 595

V. Quantitative Assay Part: Determining enzyme activity 1. Using fresh wells on your plate, add 50 µl of each sample to a well (Row C1 – 4). 2. Add 40 l of complete Z-buffer (with ME added). 3. Add 100 l of reaction solution (substrate for beta-gal) provided to you. 4. Allow reaction to proceed at 37oC for 30 to 45 min or until color develops. Your group will need to keep time on this. Start your timer when you add the reaction solution, and stop it when the plate is being read (next step). This will be your incubation time. 5. Obtain reading of the plate at 405 nm using the plate reader. Table 7.2c Results ONPG Assay at ʎ= 405 nm (Quantitative).

Cell Type

Dexamethasone OD405 reading Concentration


10-5 M




10-5 M


10-6 M


Chapter 8 Apoptosis A. Introduction Metazoan life forms require mechanisms that ensure the safe and economical disposal of superfluous and potentially harmful cells. Apoptosis is a carefully regulated mechanism of cellular suicide that does not result in loss of plasma membrane integrity or in an inflammatory response. The proper execution of apoptosis is essential for successful organogenesis during embryonic development and for maintenance of tissue homeostasis in adults. Apoptosis is a genetic programmed cellular response pathway and therefore differs from accidental cell death (necrosis) that results from cellular damages and invasions. Consequently, there are unique molecular features of apoptosis that distinguish it from other types of cell death. However different immortalized cell lines may have lost different parts of the apoptotic program and therefore respond differently to proapoptotic stimuli. Two initiation pathways are triggered by distinct events to promote apoptosis in mammals. The extrinsic pathway is activated through ligation of death receptors located in the plasma membrane that transmits the death signal into the cell‟s interior. This pathway frequently serves as a mechanism to eliminate cells during development, differentiation and tissue remodeling. The intrinsic pathway, which involves mitochondrial signaling, occurs as a response to moderate perturbation of intracellular homeostasis by various cellular stresses (e.g., hypoxia/anoxia, viral or bacterial proteins, reactive oxygen species (ROS), xenobiotics, radiation, and accumulation of misfolded proteins). The mitochondrion is an important integrator for the two pathways (although not essential for all modes of extrinsic cell death), and plays a key role in the amplification of the death signal. Cancer cells are characterized by unregulated cell proliferation and often rely heavily on glycolytic energy production. Since all “healthy” cells are equipped with the apoptotic program to prevent unregulated proliferation, one of the signs indicating that a cell mass is cancerous is their inability to undergo apoptosis in response to certain stimuli. Another use of the apoptosis pathway is in the selective killing of cells during organ development. For instance, layers of skin can be found in developing human fetuses, but they are removed by the apoptosis process as the fetus matures.

71 B. Objectives In this chapter, students will examine two different mammalian cell lines for apoptotic response under different conditions. The two cell lines that will be used will be the rat hepatoma cell lines M38 and FT02B. M38 is a cell line that maintained some proapoptotic pathways which FT02B has lost. C. Terms Trypan Blue

A dye used to detect membrane integrity of cells.


A protease that can help liberate the cells from the surface they are attached to.


A buffer containing EDTA to chelate Mg2+. Trypsin activity is inhibited by Mg2+.


A group of enzymes that are activated during apoptosis

Lipopolysaccharide (LPS)

A component of the outer cell wall of Gram-negative bacteria.

Cyclohexamide (CHx)

A chemical that inhibits protein synthesis in eukaryotes.

D. Background There are two parts to this exercise: (1) Calculating the apoptotic index and (2) detecting DNA laddering in apoptotic cells. To induce apoptosis in M38 cells, they will be treated with CHx in conjunction with varying doses of LPS. The LPS triggers a damage response, and together with the inhibition of protein synthesis conveyed by CHx, the cells will initiate apoptosis. As a control, you will investigate M38 that have not been treated by CHx, as well as FT02B cells. Apoptotic Index The apoptotic index is the percentage of apoptotic cell out of the total number of cells. In this part of the exercise, we will determine whether a cell is apoptotic or not based on their ability to take up Trypan Blue. In healthy cells, the dye will not be able to enter. In apoptotic cells where they have begun destroying the cell, the blue dye will be able to enter. The test is essentially a test of the membrane integrity of the cells. Counting the number of blue cells to be divided by the total number of cells will give us the apoptotic index. Keep in mind that this test does not distinguish between cells that died by apoptosis or necrosis.

72 To count the cells, we will first perform the staining procedure, and then do the counting on a specialized microscope slide called hemocytometer. The counting area is marked with grid lines that can be visualized under the microscope. Once the cover slip has been put onto the slide, a single layer of cells will spread out over the counting surface, and we can count the cells using the grid lines as guides. Depending on the density of your preparation, you may need to count more than 1 preparation. The number of cells per ml is: cell counts in total field x 104 (see Chapter 5.1) DNA Laddering Apoptosis is a form of programmed cell death, which differs from cell death from damage in how the process of cellular demise takes place. During apoptosis, specific enzymes (caspases) are used to execute the cellular destruction, and one of the consequences is that the DNA of apoptotic cells will display “laddering.” This is a result of cleaving the DNA at the parts that are not bound by nucleosomes. In comparison, when cells die from external damages, their DNA is degraded randomly and will not show the distinct laddering pattern.

E. Procedure

Each Group of 4

In this exercise, you will be working as groups of 4. Part I: Apoptotic Index 1. Each group of 4 will receive one 24-well plates containing 6 different wells of cells. In the following procedure, treat all the wells identically. 2. Draw off the supernatant in each well and discard it. You may use the same pipette for all wells. 3. Add 0.5 ml of versene gently, without splashing the bottom of the well where the cells are. 4. Allow the solution to sit for 1 minute. 5. Remove the versene. 6. Add 50 l of trypsin and incubate for 10 minutes at 37oC. 7. After incubation, add 50 l tissue culture medium. 8. Use a piece of parafilm to mix 25 l of trypan blue dye with 25 l of sample. NOTE: Leave the sample in the well until you are ready to count them, and prepare only the number of samples that you will use immediately. If you prepare the samples on the parafilm and let it sit out, it will evaporate and ruin your results. 9. Mix the sample with the dye by pipetting up and down twice. 10. Transfer 25 l of this sample onto the counting slide. If you don‟t get enough cells in the first 25 l, you can use the remaining 25 l. 11. Using a Nikon Alpha-2 microscope with the 10X objective, count the total number of blue and the total number of white cells. Each sample should have at least 100 total cells counted. Each student in the group should get a chance to count it.


HINT: It might be easier to train your eyes to spot the samples if you count the M38 line that has been treated with high doses of LPS first, since they will have more blue cells and they are easier to spot. Table 8.1. M38 cell response (record response of second cell line on separate sheet). Well


LPS Conc.









1 g/ml



0.01 g/ml



0.1 g/ml



1 g/ml




Apoptotic Index



Apoptotic Index

Table 8.2. Use samples for DNA laddering. Well


LPS Conc.






1 g/ml


Part IIa: DNA Laddering (60 mm dish) 1. 2. 3. 4. 5. 6. 7.

Cells for DNA laddering will be in 60 mm plates. Treat all samples the same. Pour off the culture medium in the dish. Add 400 l lysis buffer. Add 50 l protease. Mix by gently tilting the dish back and forth. Add 125 l of 4M NaCl. Mix by gently tilting the dish back and forth. Transfer everything into a clean 1.5 ml microcentrifuge tube. Label your tube. Incubate at 4oC overnight. We will keep these cells for you until next lab period.

Part IIb: DNA Laddering continued 1. Spin your sample at top speed at 4 oC for 30 min. 2. Transfer the supernatant (approximately 500 l) to a fresh tube.

74 3. Add 1 ml ethanol. Mix evenly by inverting tube several times. 4. Spin at top speed for 10 min. 5. Remove all the liquid and dry the pellet in the fume hood. 6. Resuspend the dried pellet in 20 l TE. 7. Add 1 l RNAse and incubate at 37oC for 20 min. 8. Add 1 l of 0.5M EDTA and 2 l 10X loading dye 9. Load your sample onto a 1% agarose gel.


Chapter 9 Molecular Adaptations to Temperature A. Introduction

Frequency (#)

Frequency (#)

Biological systems are complex, and temperature is one of the most important variables affecting all aspects of life. Biologists are in need of comprehensive studies assessing the impact of thermal stress on ecological systems and the species within these systems. It is a known fact that the average surface temperature of the earth has increased by approximately 0.9 °C since 1880 (NRC, 2006). We need a foundation for developing predictive models to assess the success or failure of organisms to cope with these increasing temperatures. The impact of temperature on biological systems occurs at all levels of biological organization from the molecular (e.g., the speed by which chemical reactions in the body occur) to the ecological (e.g., the distribution of species in a given habitat). A thorough understanding of temperature stress can only be obtained by investigating the consequences of elevated temperature on all levels of biological organization. Animals that are unable to maintain a constant body temperature (ectotherms) are more directly impacted by the environmental temperature than warm-blooded (homoeotherms) animals. Lake fish species are perfect models to study the effects of temperature on the physiology and the distribution of animals because these 250 Lake Coffeen species are ectothermic and survive in their natural 200 habitat within a narrow range of temperatures. The ability of a given species to cope with elevated 150 temperatures depends mostly on inherited (genetic) 100 factors, but difference in the upper limit to tolerate 50 thermal stress can vary by geographical regions for 0 the same species (genetic plasticity). These Lake Mattoon differences are thought to be the result of long-term 200 adaptations. However, the mechanisms that are 150 necessary for a species to adapt rapidly to an 100 increase in environmental temperatures are poorly 50 understood. 0

A perfect system to study the impact of thermal stress on fishes is Coffeen Lake in Montgomery County, Illinois. Coffeen Lake is a 445 ha cooling reservoir for a 945 megawatt coal-fired power Fig 9.1: Age frequency station operated by AMEREN Energy Generating distribution of blue gill in two Company. Since the 1970s, hot-water effluent from different lakes in Illinois. the plant impacts roughly 75% of the lake and water temperatures of up to 40 º C (104 ºF) have been frequently recorded (Colombo et al., unpublished data). This is roughly 25 ºC above the temperature of a „normal‟ lake in Illinois. Our preliminary data shows bluegill in Coffeen Lake survive but never reach the older ages commonly found in lakes that are not „superheated‟ (shown in figure 9.1), like Lake Mattoon. Lake Mattoon is a 424 ha reservoir located in Shelby, Cumberland, and 0




Age (Years)



76 Coles Counties with a similar bluegill similar to Coffeen Lake allowing direct comparisons to be made.

B. Objectives In this exercise, students will be introduced into scientific process by developing a testable hypothesis that investigates the molecular basis for the severely reduces lifespan of bluegill in Lake Coffeen. C. Background The impact of temperature on natural systems is of growing concern due to global climate change predictions. Current predictions suggest a 0.2 °C per decade increase at the current level of fossil fuel emissions (IPCC 2007). How this predicted change in temperature will impact species that do not maintain a constant body temperature is essential for the management of aquatic systems. Temperature can have large impacts on aquatic organisms. Often high temperature can be directly lethal. However, increased temperature can also lead to sublethal effects that impact populations over long periods of time. Bluegill (Lempomis machrochirus) provides an ideal study organism for determining the impact of temperature on fishes. The bluegill has a large spatial distribution, is found in all 48 contiguous states, and is economically important since it is the mayor game-fish in several states. Additionally, because of its importance, much of its life history parameters (growth, mortality, and lethal temperatures) have been elucidated. Assessing the molecular responses of bluegill to increasing temperature requires a model system that has an altered temperature regime. Coffeen Lake provides this opportunity with a population of bluegill that has been exposed to extremely high temperatures for more than three decades. By comparing the population in Coffeen Lake against the „normal‟ acclimated population in Mattoon Lake, we have the unique ability to assess the impact of an altered temperature regime on the physiology of freshwater fish populations. Thoroughly understanding the mechanisms by which bluegill survive with reduced longevity in the hot waters of Coffeen Lake will allow better assessments of consequences of global warming and provide novel insights into the molecular basis of aging. D. Procedure

Each Table

Design your experiment as a group after meeting with your advisor. Bring your experimental design with to our group discussion (Assignment 3, 20 points).


LAB REPORT CHECK LIST MOST IMPORTANT: ____ Does my Lab Report resemble the “before” example manuscript? Body of Text ____ Title page included and in proper format. ____ Document double spaced. ____ Clearly written; concise, terse, succinct. ____ No slang or jargon. ____ All abbreviations and acronyms spelled out upon first use. ____ No occurrences of "It can be seen that…”, “It is clear that…”, “We concluded…" or "We can see…". ____ No references to the course Instructor. ____ Proper verb tense. ____ Include headings for each section. ____ Very few, if any, quotes. ____ Not a cookbook. ____ No sentences starting with a digit. (“2 test tubes…” should be “Two test tubes…”) ____ Proper English throughout. ____ Proofread it! Scientific merit ____ References in alphabetical order and in the proper format. A minimum of ten references is used. URL’s don’t count! Preparation of Tables and Figures

78 ____ Data are presented in either a figure, a table, or in the text, and not duplicated. ____ In the text of the paper always refer to every table and figure included. ____ No unnecessary tables or figures in which the data could be replaced by text. ____ No titles on figures, each figure has a legend and all legends are grouped together on a separate page. ____ Numbers in tables have the same precision and always have a leading zero. ____ No occurrences in legends of “A graph showing…”, “A Figure of…”, etc. ____ Tables all have a table number followed by a legend. ____ Tables have appropriate and precise column headings. ____ In tables, like elements read down. ____ In tables, no columns of data that contain the same value in every cell. ____ Tables contain horizontal rules (lines) only. ____ In figures, both axes are labeled and include units. ____ No “full page” figures. ____ In figures, match the extent of the axes to the data. ____ Each figure is on a separate page. ____ No grid lines on figures unless they are specifically required to make a point. ____ All figures in black and white, no color, with white background. ____ Figures have appropriate aspect ratio (close to 3:4). ____ No figure legends on figures. ____ Figures that look the way you want them to look, not the way the software wants them to look. ____ Lanes labeled on figures of gels.


Bioinformatics 2 A. Establishing Workbench account Go to:

1. Establish account. Choose your user name and password 2. Change menus to buttons 3. Under session tools: New Session: Bioinformatics 1 B. Obtaining nucleotide sequences from NCBI: 1. Choose nucleotide 2. Search for: „Late embryogenesis abundant‟

3. Change to more and all other 4. Choose: ?


C. Working in Workbench A. Inserting nucleotide sequences and translation into proteins 1. Copy FASTA Sequence for each gene products into your Workbench Tool Folder: Nucleotides


2. Translate each into its six possible reading frames 3. Choose longest reading frame. 4. Import sequence 5. Return to Nucleic tools 6. Obtain:

B1. Investigation Phylogenetic Relationships (What is a gene family?)


1) Find CLUSTALW alignments for nucleotide and for protein 2) Paste into a WORD document 3) What are the differences in information content between phylogenetic comparison on the level of the protein sequence or nucleotide sequence? Which sequence is more “conserved” during evolution?


B2. Adding new sequence from C. elegans 1. Go to pubmed 2. Follow instuctions in B 3. Choose C. elegans 4. Copy nucleotide sequrence: AF016513.1 (Caenorhabditis elegans Ce-LEA (lea) mRNA, complete cds) 5. Use sixframe and choose longest open reading frame 6. Generate new tree

84 B3. Comparing LEA proteins with Na+/K+ATPases

1) Go back t the NCBI database and choose Protein

2) Retrieve the following sequences for Na/K+-ATPases: NP_036637 NP_653300 NP_036636 3) Copy and paste the protein sequences into your PROTEIN folder in biology Workbench. 4) Generate a rooted tree


5) How many families of proteins can be easily distinguished from each other?


Bioinformatics 3 I. The Basic Local Alignment Search Tool (BLAST) The Basic Local Alignment Search Tool (BLAST) finds regions of local similarity between sequences. The program compares nucleotide or protein sequences to sequence databases and calculates the statistical significance of matches. BLAST can be used to infer functional and evolutionary relationships between sequences as well as help identify members of gene families. You will determine identity of an unknown protein by using BLAST (Basic Local Alignment Search Tool). The tool can be found at: The sequence for the unknown protein you are investigating can be found on D2l: „TEST SEQUENCE FOR B2‟ A. Identifying an unknown protein (4pts): 1. Which type of sequence did you get? (1pt) 2. Which type of comparison should you choose? (1pts) 3. What is the identity of your protein (match: 100%)? Give at least two alternative names for the protein (1pts) 4. How is the protein activated (see ‘SUMMERY’ in best match)? (1pts)

Figure 1: The Basic Local Alignment Search Tool.


B. Judging phylogenetic relationships (4 pts): Several mathematical models can be used to investigate evolutionary relationships among genes based upon similarities and differences in their sequence. Phylogenetic trees are constructed by using computational phylogenetics methods. NCBI offers two simple methods to construct a phylogenetic tree: ‘Fast Minimum Evolution’ and ‘Neighbor Joining’. ‘Neighbor Joining’ is a clustering method used for the construction of phylogenetic trees, created by Naruya Saitou and Masatoshi Nei. The Figure below shows a proposed phylogeny for the great apes, Hominidae, taken in part from Purvis [Purvis, 1995]. The tree consists of a number of nodes (also termed vertices) and branches (also termed edges). These nodes can represent an individual, a species, or a higher grouping and are thus broadly termed taxonomical units. In this case, the terminal nodes (also called leaves or tips of the tree) represent extant species of Hominidae and are the operational taxonomical units (OTUs). The internal nodes, which here represent extinct common ancestors of the great apes, are termed hypothetical taxonomical units since they are not directly observable.

Figure 2: A proposed phylogeny of the great apes (Hominidae). Different components of the tree are marked, see text for description.

We will now investigate revolutionary relationships of other proteins to our protein. Go back to your blast result page. Under other reports choose Distance tree of results:


Figure 3: Distance tree of phylogenetic relationships.

Under „Tree method‟ choose: „Neighbor Joining‟. Under „tree form‟ choose: „rectangle‟

Figure 4: Choosing tree method and output format.

1. Which proteins is the closes relative to our protein? Hint your protein will be shown in blue as „unnamed protein product‟: (1pt) 2. What have the members in our „family branch‟ (consisting of ~8 nodes) in common? (1pt) 3. What is the second closest group of proteins (containing 2 nodes)? (1pt) 4. Is our protein closer related to the protein from rodents or the protein frogs? (1pt)

89 I. The Expert Protein Analysis System (ExPASy) proteomics server The Expert Protein Analysis System (ExPASy) proteomics server of the Swiss Institute of Bioinformatics (SIB) is dedicated to the analysis of protein sequences and structures. It contains numerous tools to analyze proteins and to judge possible functions and cellular localizations „in silicio‟. The server functions in collaboration with the European Bioinformatics Institute. ExPASy also produces the protein sequence knowledgebase, UniProtKB/Swiss-Prot, and its computer annotated supplement, UniProtKB/Trembl. Between its installation on 1 August 1993 and 5 April 2007, ExPASy

was consulted 1 billion times. The tool can be found at: Figure 5: ExPASy proteomics server.

A. You decide that you want to purify your protein via column chromatography. Since you want to choose the right columns you are interested into some basic physiochemical properties of your protein. Under tools and software choose: ‘Proteomic tools’. Since you are interested in the basic parameters choose: ProtParam. (4 pts)

Figure 6: ExPASy proteomics tools.

90 1. List the molecular weight (the weight displayed is in Dalton), the isoelectric point (pI), and the number of amino acids. Cleave off the first 44 amino acids and note the changes in molecular weight and pI. (1 pt) 2. Based on the number of amino acids and the weight of your protein what is the average weight of an amino acid in Dalton? (1pt) 3. You choose anion exchange chromatography to purify your protein. In order for your protein to bind to the positive matrix of the anion exchange column you must make sure that your protein is negatively charged. At which pH values will your protein be negatively charged? (1pts)

B. You are interested in investigating the consequences of the protein being transported to different subcellular locations. Therefore, you first want to know about the subcellular fate of your protein „in vivo’. Several prediction tools are available at ExPasy (e.g. TargetP, psort). Some of the prediction tools are designed to specifically ask about localization to a specific compartment (e.g. ChloroP, MITOPROT, Predator) whereas other approaches are more generalized (e.g. psort). These programs often compare your sequence with known sequences and give you the likelihood that your protein is directed to a ceratain compartment or excreted. However, even this software is a useful tool to investigate your protein; the final judgment whether your protein is localized to a certain compartment in vivo will have to be done in an experiment at the „wet-bench‟. However, you decide to use psort for the „in silicio’ experiment. (4 pts)

Figure 7: Psort subcellular prediction tool.

91 Under psort you choose WolF PSORT (Horton et al., 2005). Select the type of organism your sequence is derived from (Animal). Copy and paste your sequence. WoLF PSORT predicts the subcellular localization sites of proteins based on their amino acid sequences. The overall prediction accuracy of WoLF PSORT is over 80%.

Figure 8: Psort subcellular prediction tool for eukaryotic organisms.

You will be provided with a list of likely cellular localizations. As higher the number as more likely the protein will localize to that compartment. 1. What is the most likely localization of the protein? (1pt) 2. You decide to see whether the „Essentials in Cell Biology‟ textbook is correct and can help you to design a protein that is located in the nucleus of the cell. You will use the „Import into the Nucleus‟ sequence from Table 15-3 from your textbook and add the sequence to the front of the protein. Before you do the „wet-bench‟ experiment in the laboratory you will use WoLF to confirm that the protein will be directed to the nucleus. Make sure you change the amino acid code from the text book into the one letter code WoLF needs. How does this addition of amino acids change the prediction of subcellular localization? (1 pt) 3. You realize that in order to transport the protein into the nucleus you must firs remove the sequence that directs the protein to be excreted. Therefore you cleave of the first 44 amino acids and then add the „Import into Nucleus‟ sequence from your

92 textbook to the truncated protein. How does this change the localization probability for your protein? (1pt) 4. Now you express your newly designed protein into epithelia cells to confirm your experiment worked at the „wet-bench‟. Soon after you initiate expression all your cells die. You realize you made a terrible mistake. What is the mistake and why do your cells die? (1pt)

Figure 9: Key to Psort subcellular prediction.



Figure 10: Three or one letter amino acid code.



BIO3120: Cell and Molecular Biology Course Syllabus Spring 2013 Instructor: Dr. Michael A. Menze 2029 Life Sciences (217) 581-6386 [email protected] Lecture/Lab: Sec 3:

LFS 2081

8:00 – 8:50 MW

8:00 – 9:50 TR

LFS 2029

9:00 – 10:30 MW

10:00 – 11:00 TR

Office hours:

You may also find me outside my office hours in LFS 2010 or LFS 2029, but it‟s best to let me know in advance or to make an appointment. Text Book and Lab Manual: We will use the text book: Essential Cell Biology, Alberts et al., 3rd Edition, Garland Publishing. The Laboratory Manual must be purchased from the Biological Sciences Graduate Student Association in LFSA 1120. Credits: 4 semester hours. Course Objectives: A deep and thorough understanding of biology requires knowledge of the structures and functions of cellular and molecular components of living cells. This class combines lectures and practical exercises to understand, explain, and analyze the basic function of cells, cellular components, energy metabolism, propagation of heritable material, and the regulation of gene expression. Students will be able to apply the principles that are learned in this course to enhance their understanding of other aspects of Biology such as evolution, ecology, and organismal physiology. The exercise sections will introduce the students to common techniques in molecular biology and allow them to gain handson experience in these techniques. The students will also be trained in how to formulate, conduct, and report scientific experiments.

96 Course Policies: I.


Intellectual Honesty – Students are expected to abide by the intellectual honesty policy of EIU. Cheating, plagiarism, lying, or falsifying of data will not be tolerated. Respect and Diversity – EIU supports a diverse learning environment, where every student can feel respected and nurtured. Students are expected to contribute to this respectful and diverse learning environment. The classroom should be free from distractions caused by the use of cell phones (including texting), inappropriate use of laptop computers and listening devices, and offensive language.

Course mechanics: I.

The course alternates between lab exercises and lectures. Experimental work as a scientist does not always allow following a strict time line. Therefore, modifications to the course schedule may be needed as the course progresses. Appropriate announcements will be made in class for any changes. Missed laboratory work or examinations can be made up only when accompanied by an acceptable written excuse. As a general rule lectures will be given Mondays and Wednesdays and practical exercises on Tuesdays and Thursdays. Exams are given before the lab part on Tuesdays and Thursday.


Homework assignments are based on the laboratory experiments and lecture material. Participation in the laboratory is necessary to complete homework based on the laboratory experiments. A penalty will be assessed for home work turned in after the due date. Students are expected to commit additional time to these assignments as needed.

Course Grades: Lecture and Lab related exercises: I. Exams – 4, 100 points each: II. Worksheets – 10 worksheets, 15 points each: III. Lab journal – collected twice, 15 points each: IV. Lab report: V. *Group assignments – 3 :

400 points 150 points 30 points 70 points 50 points

Total possible:

700 points

(55%) (20%) (5%) (10%) (7%)

*For detailed instructions on group assignments refer to page 5 and page 58 of the lab manual. Lab quizzes are given via D2l about a week before the beginning of the next lab session. Upon completion of a lab experiment, each student should hand in the

97 associated lab worksheet in class. If you miss a lab, your worksheet for that lab will not be accepted. The worksheet will also contain a lecture related quiz section. Exams are given in class and will consist of multiple choice questions, fill in the blank, definitions, and short assay questions. Exam will be based on BOTH Lab and Lecture. Exams will be given before the lab part on Tuesdays and Thursdays. Group Assignments will be take home group exams. The maximal amount of possible points is 700 points. A total score of 90% or above assures a grade of „A‟, 80% or above „B‟, 70% or above „C‟, 60% or above „D‟. Each student will need to keep a lab journal. In the journal you will note any changes to the procedures described in the lab manual, summarize your results, and offer a brief discussion of the results. The journal will be collected twice during the semester and graded. The lab report should be in the form of a scientific publication. Please consult the laboratory manual for a detailed explanation.

Lab journal (bound notebook NOT a three ring binder): A) For each experiment the lab journal should contain one page divided into with four sections: 1. Introduction, 2. Material and Methods, 3. Results, and 4. Discussion. B) Before each experiment write an Introduction (brief summary of the experiment) into your lab journal. You will have time to work on the other sections during and after the lab. C) Lab journals will be collected on two random occasions during the semester. The journals will be graded based on completeness (5 pts), readability (5 pts), and scientific content (5 pts). D2L: This course will utilize D2l. Students who are not familiar with the software should seek training from the ITS. A tutorial on how to navigate D2l can be found here: Some of the materials you will need for completing some lab exercises will be delivered through the D2l. Students are responsible for checking the course page on D2l to keep up with the course work. I strongly suggest that you check D2l at least once a day. Lab fee: A fee of $35 will be assessed from each student enrolled on the 10th class day to help with the cost of laboratory supplies.

98 Disabilities: Persons with special needs should make those needs known to the instructor at the beginning of the course. Course Schedule: Laboratory and lecture related exercises:


Are assigned after completion of a lab experiment.


Will be given on 01/24, 02/19, 04/02, and during the finals week.

Lab report:

The lab reports is due on 04/11

Lab journal:

Collected twice unannounced

99 Appendix: I. Readings for lectures and exams*: Week 1 1

Lecture 1 2

1 1 1 2 2 3 4 4 5 5 6 6 7 8 8 9 9 11 11 12 12 13 13 14 14 15 16 16

3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30

Topping DNA Structure and Function I DNA Structure and Function II Protein basics DNA Replication I DNA Replication II DNA Repair and Recombination Transcription and RNA Transcription II and Translation of mRNA I Translation of mRNA II Protein Structure and Function I Protein Structure and Function II Protein Structure and Function -Enzymes III Membrane Structure and Function I Membrane Structure and Function II Membrane Transport I Membrane Transport II Intracellular Compartments and Transport I Intracellular Compartments and Transport II Intracellular Compartments and Transport III Cellular Signaling I Cellular Signaling II Cellular Signaling III Cell Cycle Control I Cell Cycle Control II Cytoskeleton I Cytoskeleton II Mitochondrion 1 Mitochondrion 2 Glycolysis and Fermentation TCA cycle I TCA cycle II and Bioenergetics

*Schedule and topics may changes at the discretion of the instructor.

Pages 171-189 171-189 119-121 197-210 210-221 211-221 231-241 241-251 252-260 122-142 142-156 142-156 361-385 361-385 387-421 387-421 495-526 495-526 495-526 531-568 531-568 531-568 609-624 609-624 571-606 571-606 453-475 453-475 476-494 425-436 437-452






100 2. Tentative lab and worksheet schedule*: Week 2 3 4 5


7 8 9 10 11 12 13 14 15 16

Date 01/15 TU 01/17 R 01/22 TU 01/24 R 01/29 TU 01/31 R 02/05 TU 02/07 R 02/12 TU 02/13 W 02/14 R 02/19 TU 02/21 R 02/26 TU 02/28 R 03/05 TU 03/07 R

Chapter Lab 1 Lab 2.1 Lab 2.2 Lab 2.2 Lab 2.3 Lab 2.4 Lab 2.4 2.5 B2 Lab 2.6 Lab 2.6 Lab 2.6 Lab 3 Lab 3 Lab 4.1 Lab 4.2 Lab 5.1 B3

03/19 TU 03/21 R 03/26TU 03/28 R 04/02 TU 04/04 R 04/09 TU 04/11 R 04/16 TU 04/18 R 04/23 TU 04/25 R

Lab 5.2 Lab 5.2 Lab 6 Lab 6 Lab 6 Lab 7 Lab 7 Lab 8 Lab 8 Lab 9 Lab 9 Lab 9

Topic WK/Ch Lab safety, pipetting, assignment 1 1 (1.1-2.1) Lecture plasmids, digestion Electrophoresis DNA, discuss paper (Brenner) 2 (2.2) EXAM 1, data analysis, lecture discussion Plasmid extraction, DNA quantification 3 (2.3) Digestion, bioinformatics 1, plasmid mapping Run gel, start ligation, lecture ligation Extra credit exercise „reading frames‟ 4 (2.4) E-classroom library (4450): bioinformatics 2 Transformation 1, plate transformants, lecture discussion Pick clones Transformation 3, plasmid extraction, digestion EXAM 2, PCR 1, lecture PCR, run gel 2.6 5 (2.5-3.0) PCR 2 (electrophoresis), assignment 2 Enzymatic activity: temperature 6 (4) Enzyme activity: pH Extra credit contest: counting Kc167 and Sf-21 cells 7 (B3) E-classroom library (4450): bioinformatics 3 SPRING BREAK Group Exp. 1: osmotic stress in insect cells I Group Exp. 1: osmotic stress in insect cells II Western Blot 1: protein extraction, SDS-PAGE 8 (6) Western Blot 2: antibody washes, detection EXAM 3, discussion group experiment 2 Reporter gene assays I: protein extraction Reporter gene assays II: ONPG assay Lab report ONPG analysis, discussion group experiment 2 9 (7) Apoptosis I Apoptosis II, assignment 3 Group Exp. 2: molecular adaptations to temperature I 10 (8) Group Exp. 2: molecular adaptations to temperature II

*Schedule and topics may changes at the discretion of the instructor.