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Manipulation and Expression of Recombinant DNA A Laboratory Manual
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Manipulation and Expression of Recombinant DNA A Laboratory Manual Second Edition
Susan Carson Biotechnology Education Program Department of Botany North Carolina State University
Dominique Robertson Department of Botany North Carolina State University
AMSTERDAM • BOSTON • HEIDELBERG • LONDON • NEW YORK • OXFORD PARIS • SAN DIEGO • SAN FRANCISCO • SINGAPORE • SYDNEY • TOKYO Academic Press is an imprint of Elsevier
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Cover image courtesy of Ravisha Weerasinghe. Fluorescence images of plant epidermal and root hair cells expressing Green Fluorescent Protein (GFP) fused with microtubule associated protein, MAP4 (left-hand panel) and actin binding protein, Talin (right-hand panel). These constructs provide an excellent system to monitor cytoskeletal dynamics in living cells. New evidence confirms that root knot nematodes and rhizobia produce an essentially identical response in cytoskeletal dynamics. See article by Weerasinghe, Bird, and Allen. PNAS, Feb. 22, 2005.
Elsevier Academic Press 30 Corporate Drive, Suite 400, Burlington, MA 01803, USA 525 B Street, Suite 1900, San Diego, California 92101-4495, USA 84 Theobald’s Road, London WCIX 8RR, UK This book is printed on acid-free paper. ⬁ Copyright © 2006, Elsevier Inc. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher. Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone: (+44) 1865 843830, fax: (+44) 1865 853333, E-mail:
[email protected]. You may also complete your request on-line via the Elsevier homepage (http://elsevier.com), by selecting “Support & Contact” then “Copyright and Permissions” and then “Obtaining Permissions.” Library of Congress Cataloging-in-Publication Data Application submitted. British Library Cataloguing in Publication Data A catalogue record for this book is available from the British Library ISBN 13: 978-0-12-088418-6 ISBN 10: 0-12-088418-6 For all information on all Elsevier Academic Press publications visit our Web site at www.books.elsevier.com.
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Contents xi
Preface
Acknowledgments
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Note to Instructors
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Instrumentation
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Nomenclature
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INTRODUCTION
Conceptual Outline for Experiments I. II. III. IV.
EXPERIMENTAL PROCEDURES 1 LABORATORY SAFETY 2 GENERAL OPERATING PROCEDURES EMERGENCY CONTACT INFORMATION
1
4 4
PA R T I
Manipulation of DNA LAB SESSION 1
Getting Oriented; Practicing with Micropipettes I. STATION CHECKLIST 7 II. MICROPIPETTING 9 III. LABORATORY EXERCISE
7
11 11
A.
Preparing BSA Dilutions
B.
Performing a Nitrocellulose Spot Test
12
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LAB SESSION 2
13
Large-Scale Purification of Plasmid DNA I. INTRODUCTION 13 II. LABORATORY EXERCISES
14
A.
Alkaline Lysis and Anion Exchange Chromatograhy
B.
DNA Quantification
14
17
LAB SESSION 3
Preparation of Expression Vector DNA (pET-41a(+), a GST Fusion Protein Vector) 19 I. INTRODUCTION 19 II. LABORATORY EXERCISES
23
A. Restriction Digestion of Vector (pET41a) Restriction Enzyme Digestions 23
24
B.
Agarose Gel Electrophoresis
C.
Cleaning DNA Using a Spin Column
26
LAB SESSION 4
Preparation of Insert DNA (egfp) I. INTRODUCTION 29 II. LABORATORY EXERCISES
29
30
A.
Restriction Digestion of pEGFP-N1
B.
Isolation of egfp DNA from Agarose
30 30
LAB SESSION 5
Preparation of Transformation-Competent Cells and Control Transformation 35 I. INTRODUCTION 35 II. LABORATORY EXERCISES
35
A. Preparation of Chemically Competent Cells by Calcium Chloride Treatment 35 B.
Transformation Control
36
LAB SESSION 6
DNA Ligation and Transformation of Escherichia coli I. INTRODUCTION 39 II. LABORATORY EXERCISES
41 41
A.
Ligations and Ligation Controls
B.
Divalent Cation-Mediated Transformation
C.
Electrophoresis of Ligation Reactions
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Screening Transformants LAB SESSION 7
47
Colony Hybridizations LAB SESSION 7A
Interim Laboratory Session I. INTRODUCTION 49 II. LABORATORY EXERCISES A.
Counting Transformants
B.
Replica Plating
49 49 49
50
LAB SESSION 7B
Colony Hybridization: DNA Probe I. INTRODUCTION 53 II. LABORATORY EXERCISES
53
57 57
A.
Colony Hybridization with an egfp DNA Probe: Part 1
B.
Labeling of DNA Probe by PCR Using Digoxigenin-11-dUTP
57
LAB SESSION 7C
Colony Hybridization: Monoclonal Antibody Probe I. INTRODUCTION 59 II. LABORATORY EXERCISE
59
60
A. Colony Hybridization with an a-GFP Monoclonal Antibody Probe: Part 1
60
LAB SESSION 8
Completion of Colony Hybridization with DNA Probe I. INTRODUCTION 63 II. LABORATORY EXERCISE A.
63
63
Colony Hybridization with an egfp DNA Probe: Part 2
63
LAB SESSION 9
Characterization of Recombinant Clones
65
LAB SESSION 9A
Completion of Colony Hybridization with mAB Probe I. INTRODUCTION 67 II. LABORATORY EXERCISE
67
67
A. Colony Hybridization with an α-GFP monoclonal Antibody Probe: Part 2
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LAB SESSION 9B
PCR Screening
69
I. INTRODUCTION 69 II. LABORATORY EXERCISE A.
71
Polymerase Chain Reaction Screen for Recombinant Clones: Part 1
71
LAB SESSION 9C
Visualization of Green Fluorescent Protein: Part 1 I. INTRODUCTION 73 II. LABORATORY EXERCISE A.
73
73
Green Fluorescence Assay and Preparation of a Fresh Master Plate
73
LAB SESSION 10
Further Characterization of Recombinant Clones
75
LAB SESSION 10A:
Interim Laboratory Session I. LABORATORY EXERCISE A.
77 77
Inoculate Cultures for Minipreps
77
LAB SESSION 10B
Analysis of PCR Screen Results I. INTRODUCTION 79 II. LABORATORY EXERCISE A.
79
79
Gel Electrophoresis and Analysis of PCR Samples
79
LAB SESSION 10C
Isolation and Characterization of Miniprep DNA from Potential Transformants (Restriction Analysis of Putative Transformants) I. INTRODUCTION 81 II. LABORATORY EXERCISES
81
82
A.
Alkaline Lysis and Ethanol Precipitation of Miniprep DNA
B.
Restriction Enzyme Analysis of Miniprep DNA
82
83
LAB SESSION 10D
Visualization of Green Fluorescent Protein: Part 2 I. INTRODUCTION 85 II. LABORATORY EXERCISE
85
85
A. Visualization of Clones Expressing the Enhanced Green Fluorescent Protein on IPTG Plates 85
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Expression, Detection, and Purification of Recombinant Proteins from Bacteria LAB SESSION 11
Expression of Fusion Protein from Positive Clones and Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE) and Immunological Analysis (Western Blot): Part 1 91 LAB SESSION 11A
Interim Laboratory Session I. LABORATORY EXERCISE A.
93 93 93
Inoculate Cultures for SDS-PAGE
LAB SESSION 11B
Expression of Fusion Protein from Positive Clones and Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE) and Immunological Analysis (Western Blot): Part 1 95 I. INTRODUCTION 95 II. LABORATORY EXERCISE A.
96
SDS-Polyacrylamide Gel Electrophoresis and Western Blot: Part 1
96
LAB SESSION 12
Expression of Fusion Protein from Positive Clones and Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE) and Immunological Analysis (Western Blot): Part 2 101 I. INTRODUCTION 101 II. LABORATORY EXERCISES
102
A.
SDS-Polyacrylamide Gel Electrophoresis and Western Blot: Part 2
B.
Replica Plate Positive Clone
103
LAB SESSION 13
Extraction of Recombinant Protein from Escherichia coli Using a Glutathione Affinity Column 105 LAB SESSION 13A
Interim Laboratory Session I. LABORATORY EXERCISE A.
107 107
Inoculate Cultures for Protein Purification
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LAB SESSION 13B
Extraction of Recombinant Protein from Escherichia coli Using a Glutathione Affinity Column 109 I. INTRODUCTION 109 II. LABORATORY EXERCISES
112
A. Growing Bacterial Suspension Cultures for Fusion Protein Purification B.
Harvesting IPTG-Induced Cultures
C.
Breaking Open Bacterial Cells
112 112
D. Removing Insoluble Debris from the Crude Homogenate E.
Purifying Protein by Affinity Chromatography
113
LAB SESSION 14
Analysis of Purification Fractions I. INTRODUCTION 115 II. LABORATORY EXERCISES A.
115
117
SDS-PAGE of purified Fusion Protein
117
B. Bradford Protein Concentration Determination Assay of Purification Fractions 118
APPENDIXES 1. EQUIPMENT 121 2. PREP LIST 123 3. MAKING SENSE OF ORIENTATION INDEX
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143
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Preface he manipulation and expression of recombinant DNA is deceptively simple: cut and paste DNA together and new proteins can be made. But to really understand the power of biotechnology, there is no adequate substitute for sequential, wet lab experiments that lead students through the process from isolating DNA to making recombinant protein. Eight years have passed since the first edition of this manual was published. The second edition provides updated experiments and techniques but the core principles remain the same: combining nucleic acids from different organisms to produce a recombinant product. The experiments described in this manual continue to provide a solid conceptual basis for understanding the power and promise of biotechnology, even as the field evolves. In our experience, students from many different disciplines, including life sciences, agriculture and veterinary medicine, chemical engineering, and physics have benefited from these experiments. Each of these disciplines uses biotechnology differently but the principles are the same. One of the most rewarding parts of working with students is watching them transition from knowing about biotechnology to “Yes, I see the applications of biotechnology.” Once the power of recombining nucleic acids to produce novel structures is demonstrated, advanced biotechnologies in health care, agriculture, and nanotechnology become apparent. When this manual was first written (1997), the human genome was not yet sequenced and products developed through biotechnology were just beginning to be marketed. Now, most of the corn, soybean, and cotton grown in the United States have been genetically altered to the benefit of farmers. Consumer benefits are in development and plants have been modified to detoxify methyl mercury, detect gunpowder, and express vaccines. Our knowledge of the biological basis of human genetic disease has skyrocketed, and noninvasive methods for treating cancer and other diseases are beginning to emerge. There is no longer any doubt that the quality of life can be improved by biotechnology. Mistakes have been made, such as the contamination of tacos meant for human consumption by transgenic maize approved only for animal feed in 2001. However, the political and social consequences of such mistakes are severe and Aventis no longer produces transgenic plants. While the caveat remains that any technology can be used for good or bad, we must ask if we can afford not to use biotechnology to address quality of life issues. The continued safeguarding of our food supply and environment will require scientists, corporations, and governments to work together to formulate policies that promote technological innovations. Long-term initiatives that address global demands, restoration of natural resources, and international cooperation can and should include biotechnology. It is our hope that years from now, biotechnology will be seen as a highly refined tool to fight disease, promote a healthy, ecologically robust planet, and ensure quality of life for everyone.
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Acknowledgments e would like to thank the many people who contributed to this manual. Melissa Cox and Anna Douglas were instrumental in piloting the experiments to make sure they work as they should. Thanks also to Lisa Lyford, Melissa Cox, Beth Rueschhoff, Jennifer Modliszewski, and Wenheng Zhang for comments on the manuscript. The NCSU undergraduate and graduate students in BIT 360 and BIT 810 who willingly acted as guinea pigs to test this new course for use in the classroom were wonderful and enthusiastic. We also thank the North Carolina State University Colleges of Agriculture and Life Sciences, Engineering, Veterinary Medicine, Physical and Mathematical Sciences, and Natural Resources. The support from these colleges for the Biotechnology Program led to the development of this manual.
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Note to Instructors hese laboratory exercises were developed in the context of the curriculum offered by the North Carolina State University Biotechnology Program (http://www.ncsu.edu/biotechnology). Students take the Manipulation and Expression of Recombinant DNA course as a prerequisite to more specialized laboratory courses, including Fermentation of Recombinant Microorganisms, Protein Purification, Animal Cell Culture, Microarray Technology, PCR and DNA Fingerprinting, RNA Purification and Analysis, and others. The laboratories in this course prepare students well both for these specialized courses, and for independent research in a molecular biology laboratory, both at the undergraduate and graduate student levels. Our Manipulation and Expression of Recombinant DNA course is a 4-credit lecture/lab course. We meet for 2 hours once per week for lecture, and allot 5 hours for one lab period per week. We recommend this schedule because for several of the labs, students must inoculate cultures or perform other short activities prior to their lab day. It works out well for them to do so at the end of their lecture period (these activities are listed as “interim laboratory sessions” in the Contents). The majority of the laboratories do not require the full 5 hours, but a few of them do. Additionally, there are a few labs for which incubation times are simply too long for them to be included in the exercises. In these cases, the steps are included in the protocols with a note that the instructor will perform that particular part of the experiment for the class (for example, the induction of the fusion protein using IPTG, necessary for several laboratories, takes 3 hours). For this reason, we recommend offering the laboratory as an afternoon course so that the instructor can begin the incubations in the morning, rather than in the middle of the night. If your semester is too short to accommodate all 14 lab sessions, the exercise described in Lab Session 5 can be omitted. It is included as an inexpensive alternative to commercially available competent cells. Frozen competent cells can be purchased from a variety of vendors. Alternatively, competent cells can be prepared by the prep staff in bulk and stored at −80˚C. The commercial cells have the advantage that they often have a higher transformation efficiency than the home-made cells. In this manual we often refer to “lab stations.” This course was designed for students to work in pairs. Each pair of students is assigned a particular bench that they use every week. We number the benches and refer to them as lab stations. Students label all of their experiments, cultures, and so forth with their station number rather than their initials. All antibodies and plasmids described in this manual are available commercially, with the exception of the positive control, pAD1. A small quantity of this plasmid is available at no cost (other than shipping) from Dr. Sue Carson at NCSU. Contact Dr. Carson at
[email protected], and include in the subject heading
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Note to Instructors
“pAD1 request.” Appendix 2 lists company and catalog numbers of plasmids and E. coli host strains used in this manual.
Instrumentation Certain lab sessions provide detailed instructions for using a particular brand-name apparatus. Similar equipment from other vendors can be substituted. Appropriate instrument-specific instructions should be substituted to minimize confusion. This is especially true of the DNA agarose gel electorphoresis units, the protein polyacrylamide gel electrophoresis units, the transfer apparatus for Western blotting, and the sonicators. Essential laboratory equipment is listed in Appendix 1.
Nomenclature In the literature, the nomenclature for the abbreviations of the enhanced green fluorescent protein gene and its gene product has been inconsistent at best, and downright confusing at worst. In this publication, we will use “egfp” to refer to the gene and “EGFP” to refer to the gene product. Likewise, we will use “gst” for the glutathione-S-transferase gene and “GST” for its gene product. Bacterial genes discussed in this book will use standard bacterial nomenclature with the gene name lowercase and italicized, and the gene product with a capitalized first letter and not italicized. For example, the gene for the lac repressor is “lacI” and its gene product is “LacI.”
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CONCEPTUAL OUTLINE FOR EXPERIMENTS oal: . Make a fusion protein by splicing genes from two organisms: one from Escherichia coli (gst) and an enhanced gene derived from the green fluorescent jellyfish Aequoria victoria (egfp). Expression of the fused gene will produce a single protein in bacteria. The E. coli part of the fusion protein will be used as a tag to purify the fusion protein. The A. victoria portion of the fusion protein can then be characterized or used for antibody production.
G
I. EXPERIMENTAL PROCEDURES (See Figure 1 for a diagrammatic representation.) ●
●
● ●
●
● ●
● ●
●
●
Isolate plasmid DNA using large-scale cultures of bacteria containing either the cloned A. victoria gene or the E. coli expression vector. Use restriction enzymes to cut the vector (containing the E. coli gst gene) and the insert (A. victoria egfp DNA). Use DNA ligase to “paste” the vector and insert DNA together. Introduce the ligated DNA into E. coli. Identify bacterial transformants having A. victoria DNA inserts by colony hybridization using purified, labeled A. victoria DNA as a probe. Identify bacterial transformants that correctly express the A. victoria DNA in a fusion protein by immunoassays. Confirm positive clones by Polymerase Chain Reaction analysis. Isolate DNA from bacterial clones positive for both the DNA and monoclonal antibody probes. Digest DNA with restriction enzymes to further confirm the presence of the A. victoria gene and verify that only one copy was inserted. Perform final confirmation of egfp-positive clones by fluorescence. Use sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) of bacterial cell preps to determine the level of expression of the fusion protein in different transformants. (This is a pilot experiment for large-scale isolation of fusion protein.) Transfer protein from SDS-PAGE onto nitrocellulose membrane to perform Western blot analysis to confirm the stability of the fusion protein. Induce a large-scale culture of the transformed bacteria with isopropyl-β-Dthiogalactopyranoside (IPTG) to make large amounts of the fusion protein.
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Purify the fusion protein on a substrate affinity column. Perform protein quantification of eluted fractions. Use SDS-PAGE of intermediates in the purification procedure and final eluates to check for purity and degradation. T7 terminator NotI NcoI gst
NcoI NcoI
kan
egfp pEGFP-N1 4733 bp
pET-41a(+) 5933 bp
NotI
T7 promoter lac operator
NcoI lac I
kan NcoI source of egfp DNA
ige
st
expression vector
No
tI d
NcoI
Nc
oI/
NotI egfp 713 bp fragment
NcoI fusion gst
egfp NotI T7 terminator
kan
T7 promoter lac operator p? 6584 bp lacI
your clone!
Fig. 1 Experimental procedure diagram
II. LABORATORY SAFETY Hazards that you may be exposed to during the course of the laboratory exercises include working with toxic compounds and ultraviolet (UV) irradiation. Special precautions must be taken when working with recombinant DNA. To ensure the safety and well-being of students and support staff, the following rules will be strictly enforced. A reckless attitude about the use of equipment or the safety of others will cause you to be dropped from the course.
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The following rules must be observed at all times in the laboratory: 1. No smoking, drinking, or eating is allowed in the laboratory. Never store food in laboratory freezers or refrigerators. 2. You are responsible for providing a lab coat. Wear your lab coat, gloves, and safety glasses when working in the fume hood or in the presence of dangerous or potentially dangerous substances. No sandals or other open-toed shoes are allowed in the laboratory at any time. 3. Always wear gloves when working with ethidium bromide, phenol, or any hazardous or potentially hazardous substance. Remove your gloves before leaving the laboratory and wash the outsides of your gloves frequently to prevent contamination of equipment, etc., with caustic agents. 4. Long hair must be tied back at all times, and avoid loose-fitting clothing to avoid hazards associated with open flames, sterile cultures, and hazardous chemicals. 5. Dispose of microorganisms, including the tubes used for their growth, in orange bags marked “BIOHAZARD” for autoclaving. Liquid medium and plastic pipettes used to transfer microorganisms will be collected in specially marked flasks containing bleach. 6. Dispose of glass only in properly marked blue boxes designated for glass disposal. This includes glass culture tubes, pipettes, and broken glass. Do not put glass or any sharp object in the orange autoclavable bags marked BIOHAZARD. 7. Keep your lab bench free of unnecessary clutter. Use cabinets and drawers for storing personal items and extra supplies, not for food. At the end of the day, your bench should be clean. Micropipettes and gel rigs have been known to wander from their home station. The best way to ensure that your equipment is there the next day is to store it out of sight when you leave. 8. Wear ear protection when working with the sonicator. 9. Always wear a UV-protective full-face shield when using the transilluminator. Your safety glasses are not UV protective. Do not try to analyze your gel on the transilluminator. Take a picture and analyze the picture. If you need to excise a band from a gel on the transilluminator, make sure you are wearing a UV-protective full-face shield. 10. Wash your hands thoroughly before you leave the laboratory. 11. Spills should be cleaned up immediately. Make sure you have an adequate supply of paper towels at your station. If you need more, see an instructor. Spill control pillows are located on top of the yellow cabinet for flammable chemicals. Notify an instructor if you spill a chemical that is marked “poison” or “caustic.” If you spill liquid containing live microorganisms, notify an instructor and pour disinfectant on the spill. 12. Immediately report all accidents such as spills, cuts, burns, or other injuries to an instructor. 13. Know the location of the fire extinguisher, eye wash station, emergency shower, and emergency exits. 14. If you have trouble with a power supply or the leads to a gel, report it to an instructor. If you see someone receiving an electrical shock, use a nonconducting object, such as a plastic beaker, to break the circuit or you may receive the shock as well. 15. Leave all laboratory facilities and equipment in good condition at the end of the class. Before leaving the laboratory, check to make sure that all electrical equipment is turned off and that the gas to the Bunsen burner is turned off. 16. No pets are allowed in the laboratory. 17. Dispose of hazardous chemicals, such as chloroform, methanol, and ethidium bromide, only in designated containers. Do not pour them down the sink. Each hazardous chemical will have its own waste container. If in doubt, ask an instructor before dumping questionable reagents down the sink.
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18. An up-to-date immunization against tetanus is strongly recommended. 19. For a tutorial on general laboratory safety, visit the website http://www. ncsu.edu/project/ungradreshhmi/evaluationModule/login.php (login is free).
III. GENERAL OPERATING PROCEDURES ●
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●
●
Reagents—Aliquoting reagents and supplies for everyone in the course at the same time is difficult. Your patience and cooperation are greatly appreciated. If you run out of a reagent or enzyme and cannot find it on a lab cart, please ask a technician or TA for more. Enzymes—Enzymes are very expensive and can be ruined by prolonged exposure to room temperature or by contamination. Always use a fresh, sterile pipet tip for each enzyme. Keep the enzyme on ice; never put it in a microcentrifuge rack at room temperature. Always add enzymes to your reactions as the last component; addition of enzyme to unbuffered solutions will kill its activity. Pipette tips—You will be provided with five racks for pipette tips: two for yellow tips, two for blue tips, and one for white (P-10) tips. You are responsible for filling these racks. Use gloves to handle the tips. A technician will autoclave these tips for you. Label your boxes with your group number and place them on the lab cart containing supplies for autoclaving. Repairs—A list of supplies and equipment at your station will be made availableto each pair of students at the first lab session. You are responsible for the equipment at your station. If you encounter any difficulties with operation, please ask for assistance. If equipment is damaged in any way, please report it so that we can get it repaired. Barring willful destructiveness, you will not have to pay for repairs. Equipment—Please make sure that you understand the proper use of this equipment before attempting to operate it. If in doubt, ask an instructor, not another student. In addition to costly repairs, improper use of this equipment can be very dangerous.
Remember: A reckless attitude about the use of equipment or the safety of others will cause you to be dropped from the course.
IV. EMERGENCY CONTACT INFORMATION At the back of this manual you will find a contact form, which you should complete and turn in to your instructor. The information on this form will be used in case of emergency.
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PA R T
I Manipulation of DNA he goal of these laboratory exercises is to fuse a bacterial gene with a jellyfish gene and to express a single protein from this hybrid DNA sequence. Why would you want to do this? Molecular shuffling of genetic sequences, or gene cloning, is a powerful tool for understanding biological processes. Using basic tools developed in Escherichia coli, we can ask questions about other, more complicated organisms. Scientists have exploited E. coli both as a workhorse for producing DNA and as a source of well-characterized sequences to direct transcription and translation of foreign DNA into protein. With genetic sequence information being produced at the rate of megabases per day, the limiting factor is not the DNA analysis (although that is a challenge!) but our understanding of the function of the products of these sequences; what do the proteins actually do? Additionally, in terms of practical biotechnology applications, it can be a huge advantage to clone the gene encoding a difficult-to-purify protein into E. coli so that the purification process can be accomplished less expensively and to a greater degree of purity (and often more ethically, especially if a human gene is involved). Among the first recombinant proteins to be produced and marketed was human insulin in the early 1980s, which has been invaluable to countless diabetics. The basic tools you will learn in this class will enable you to clone, express, and purify recombinant proteins. They will enable you to begin to probe the function of any protein for which a gene has been identified, and will give you the conceptual background needed for tackling more advanced techniques. The gene we will be cloning and expressing is egfp (the gene encoding the enhanced green fluorescent protein). The green fluorescent protein (gfp) is a naturally occurring protein found in a species of green fluorescent jellyfish called Aequoria victoria. The difference between the fluorescence of the green fluorescent protein (GFP) and the enhanced green fluorescent protein (EGFP) is that EGFP emits 35x the fluorescence (at 510 nm) of GFP when excited with ultraviolet or blue light (395 nm, with a minor peak at 470 nm): EGFP is much brighter. The increased fluorescence was achieved by making mutations in the nucleic acid sequence that resulted in a small change in the amino acid composition within the chromophore region of the protein: Ser65 -> Thr, and Phe64 -> Leu (Yang et al., 1996). The gene encoding the green fluorescent protein (and its variants, including the enhanced green fluorescent protein) is widely used in molecular biological studies as a reporter gene or marker. A reporter gene is a gene that is used to track protein expression. It must have phenotypic expression that is easy to monitor and can be used to study promoter activity in different environmental conditions, different tissues, or different developmental stages. Recombinant DNA constructs are made in which the reporter gene is fused to a promoter region of interest and the construct is transformed or transfected into a host cell or organism.
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REFERENCES Yang, T, L Cheng, and SR Kain. 1996. Optimized codon usage and chromophore mutations provide enhanced sensitivity with the green fluorescent protein. Nucl. Acids Res. 24 (22), 4592–4594.
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LAB SESSION
1 Getting Oriented; Practicing with Micropipettes oal: Starting next week, you will be working on a laboratory project that will build throughout the entire semester. Before embarking on that journey, it is important to familiarize yourself with your lab space and to master the use of the workhorses of the molecular biology lab: the micropipettes. If your instructor has not given a safety orientation yet, he or she will do so today.
G I. STATION CHECKLIST
It is important to familiarize yourself with the work environment and laboratory equipment before beginning experiments. If the laboratory space in which you are working is shared by other laboratory sections at different times, much of the equipment can be shared. There are certain items, however, such as buffers and sterile disposable items that should not be shared between lab groups. Take a moment to go through your bench, shelves, and drawers to identify equipment and reagents. Use the station checklist on the following page and notify your instructor if anything is missing from your station. Items that are indicated as “per group” should not be shared between different sets of students on different lab days. Label these items with your initials, lab day, and station number. Other items should have the station number only.
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Part I. Manipulation of DNA
Station Checklist Station Number Name _________________________
Name ____________________________
One power supply box One horizontal DNA minigel apparatus for agarose gels One P10 micropipette One P20 micropipette One P200 micropipette One P1000 micropipette One box 1000-µl sterile tips per group One box 200-µl sterile tips per group One box 10-µl sterile tips per group One ice bucket (or cooler or styrofoam box for ice) One box Kimwipes (Kimberly-Clark, Roswell, GA) One 15 ml and one 50 ml styrofoam test tube rack One pack sterile snap cap tubes (17 × 100 mm) for overnight bacterial cultures One test tube rack for snap cap tubes One autoclaved container of 1.7-ml microcentrifuge tubes in a 1-liter tricorner beaker per group Two microcentrifuge tube racks One pack disposable 10-ml pipettes One plastic (or electric) pipette pump One 50-ml graduated cylinder One 500-ml graduated cylinder One 2-liter polypropylene beaker Two 1-liter polypropylene bottles, one for distilled water and one for 1X TBE buffer per group One 250- or 500-ml Pyrex orange-capped bottle for melting agarose per group One thermal glove for handling microwaved agarose Labeling tape Permanent ink marker (Sharpie) One plastic squeeze bottle for 70% ethanol One plastic squeeze bottle for distilled water One ring-stand with clamp One pair blunt-ended forceps Two safety eye glasses or goggles (for radiation and chemicals; not for UV light protection) Two lab coats One cardboard freezer box per group Protein polyacrylamide mini gel electrophoresis unit (every other station if the unit handles 2 gels) Protein mini-transblot assembly (every other station if the unit handles 2 gels) Vortex mixer Microcentrifuge Bunsen burner Heat block Timer Parafilm
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II. MICROPIPETTING Micropipettes are the tools used to measure the very small volumes of liquid typically necessary when performing molecular manipulations. We will use 4 different micropipettes in this course. Each micropipette is accurate to measure a defined range of volume, as shown in the following table (Table 1.1). TABLE 1.1 Volume ranges of micropipettes Micropipette P10 P20 P200 P1000
Volume Range 0.5-10 µl 2-20 µl 20-200 µl 200-1000 µl
Setting the micropipettes to the desired volume can be a little tricky at first. Also, it is common for beginners to confuse the P20 and P200 since they typically use the same pipette tips; therefore, remember to check which micropipette you are using before drawing in solution. Many students accidentally measure 20 µl instead of 2 µl or vice versa because of such mix-ups. Use the following figure (Figure 1.1) and the instructions below to help with setting the micropipettes until you are confident to set them on your own. Follow the instructions below for using the micropipettes. 1. Set the desired volume by holding the pipette in one hand and rotating the dials with the other hand. Do not dial past the lower limit 000 or the upper limit (shown on the pipette; either 10, 20, 200, or 1000). Familiarize yourself with these settings. 2. Attach a tip to the end of the micropipette. To ensure an adequate seal, press the tip on with a slight twist. 3. Depress the plunger to the first stop. This part of the stroke displaces a volume of air corresponding to that indicated on the dial. 4. Immerse the tip to a depth of 2–5 mm into the liquid to be withdrawn. Immersing the tip to deeper levels will cause liquid to adhere to the outside of the tip, causing errors in measurement. 5. Allow the plunger to return slowly to its original position. If the plunger snaps back, aerosols will form, contaminating the barrel of the micropipette and your solution. 6. Wait 1 sec before removing the tip from the solution, to allow the introduced liquid to enter the pipette tip fully. A too-quick removal of the tip from the solution may result in air occupying some of the calibrated volume. Check to make sure that there are no air bubbles and that the amount of liquid corresponds to the desired amount. Develop an eye for 1-µl volumes, as these are the hardest to pipette. 7. Place the tip against the side wall of the receiving vessel near the liquid interface or the bottom of the vessel. Slowly dispel the contents by depressing the plunger until the first stop. Remaining liquid can be dispelled by depressing the plunger to the second stop. Withdraw the tip from the solution and return the plunger to its original position. Check to make sure that no liquid remains in the tip. If there is a bead of liquid, reintroduce liquid from the receiving vessel to capture the bead and slowly expel the contents. 8. Discard the tip by pressing the ejector button.
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Part I. Manipulation of DNA
Micropipette settings P1000 for 200-1000 µl aliquots 1
0
0
0
5
2
0
0
0
1 ml=1000 µl
0.5 ml=500 µl
0.2 ml=200 µl
P200 for 20-200 µl aliquots
2
1
0
0
5
3
0
0
2
0.2 ml=200 µl
0.15 ml=150 µl
0.032 ml=32 µl
P20 for 2-20 µl aliquots
2
1
0
0
5
2
0
5
5
0.02 ml=20 µl
0.0155 ml=15.5 µl
0.0025 ml=2.5 µl
P10 for 0.5-10 µl aliquots
1
0
0
0
5
0
0
5
5
0.01 ml=10 µl
0.0055 ml=5.5 µl
0.0005 ml=0.5 µl
Fig. 1.1 Micropipette settings
9. Always use a new pipette tip when pipetting enzymes, or the stock solutions may become contaminated. If you accidentally contaminate an enzyme solution, tell an instructor. Always use a new pipette tip for critical volumes, as in a dilution series, because as much as 10% of the volume may stay within the tip after delivery. 10. Working with tiny volumes requires patience and accuracy. The best way to deliver a 1-µl volume is to pick up the receiving tube and make sure that a 1-µl bead is formed on the side of the tube after delivery. In the case of enzymes, schlieren rings should be visible from the glycerol–water interface if the enzyme is dispelled directly into the solution.
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Lab Session 1
Calibration of Micropipettes Water has a density of approximately 1 g/ml at 25˚C. ●
●
●
To calibrate a 1- to 20-µl micropipette, measure the weight of 10 and 20 µl of water in a tared weigh boat on an analytical balance. For best results, average three readings at each setting. To calibrate a 20- to 200-µl micropipette, measure the weight of 100 and 200 µl of water. To calibrate a 100- to 1000-µl micropipette, measure the weight of 500 and 1000 µl of water.
If your micropipette is consistently off by more than 5%, notify an instructor. Do not tamper with the adjustment.
III. LABORATORY EXERCISE: BSA SERIAL DILUTIONS AND NITROCELLULOSE SPOT TEST The purpose of this short exercise is to get used to your lab stations and practice using the micropipettes (and test your technique). Each student will perform serial dilutions of the protein bovine serum albumin (BSA) and then compare their results against their lab partner’s results using a visualization technique that uses a proteinbinding dye. Note: While the other laboratory exercises for this course will build on each other, this one will not.
A. Preparing BSA Dilutions You will be given a tube with 15 µl of a 1-mg/ml BSA solution. Prepare a dilution series of BSA standards in 5 tubes (labeled 1 through 5) according to the scheme outlined in Table 1.2 and Figure 1.2. Attach a new pipet tip to the micropipette each time to make the dilutions. Each lab partner should do a set of dilutions. TABLE 1.2 Serial dilution scheme Tube
Dilution
Protein Concentration
12.5 µl of stock (1 mg/ml) + 37.5 µl dH20 25 µl from tube 1 + 25 µl dH20 25 µl from tube 2 + 25 µl dH20 25 µl from tube 3+ 25 µl dH20 25 µl from tube 4 + 25 µl dH20
1 2 3 4 5
250 µg/ml 125 µg /ml 63 µg /ml 31 µg /ml 16 µg /ml
Serial Dilutions for BSA 12.5 ul
BSA stock 1 mg/ml
1
25 ul
25 ul
25 ul
25 ul
37.5 ul H2O
25 ul H2O
25 ul H2O
25 ul H2O
2
3
Fig. 1.2 Serial dilution scheme diagram
4
25 ul H2O
5
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B. Performing a Nitrocellulose Spot Test Amido black is a stain that quantitatively binds protein. We will use a micropipette to deliver small amounts of the BSA serial dilutions to the nitrocellulose and then stain the nitrocellulose. This will enable you to visualize the relative protein amounts in each sample and provide visual feedback on your pipetting/dilution technique. 1. Obtain a piece of nitrocellulose (always wear gloves when handling nitrocellulose). Place the nitrocellulose on a piece of 3MM paper (Whatman, Clifton, NJ) at your station. You will share the piece of nitrocellulose with your partner. 2. Spot 2 µ1 aliquots of distilled H2O (control) and each of the BSA dilutions. One partner should spot the top row with his/her samples, and the other partner should spot a row below. Spotting of the 2 µl is best done by holding the pipette tip just above the paper. Expel liquid such that a drop forms on the end of the tip. Touch the drop to the paper and the liquid will be drawn into the paper by capillary action. CAUTION: Make certain you leave enough room between each addition such that the spots do not touch each other. 3. Place the nitrocellulose on a piece of 3MM paper and allow to air dry. 4. After the spots have dried completely, stain by placing in a tray (a square Petri dish works well for this purpose) and covering with amido black staining solution. Allow to stain for 1–2 min, with gentle shaking. 5. Pour off the stain (back into original bottle—this can be reused) and cover with methanol-acetic acid destaining solution and shake gently. Change once after 5 minutes and shake gently until background is white. 6. Place the nitrocellulose on 3MM paper to dry. Compare the intensities of each spot. Do the intensities of your spots match your lab partner’s? Does each spot appear to be half as intense as the last? If not, you need to practice your micropipetting technique.
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LAB SESSION
2 Large-Scale Purification of Plasmid DNA oal: Today you will isolate plasmid DNA to use for cloning. Two plasmids will be purified. The first is pET41a, the expression vector. The second is pEGFP-N1, the source for the gene to be cloned (egfp). One half of the class will purify pET41a and the other half will purify pEGFP-N1. You will perform the plasmid purification using the QIAfilter Plasmid Maxi Kit. This protocol starts with an alkaline lysis procedure to break open the cells and separate the plasmid DNA from chromosomal DNA, and is followed by anion-exchange chromatography for further purification from cellular proteins and other cellular debris. We will then quantify the DNA.
G I. INTRODUCTION
Escherichia coli is the workhorse of molecular biology, serving as a factory for the synthesis of large amounts of cloned DNA. Today you will isolate plasmid DNA from E. coli for in vitro manipulation. Plasmid DNA is cloned in bacteria; that is, identical copies are made and propagated in bacteria. These bacterial cells are a complex mixture of plasmid DNA, chromosomal DNA, proteins, membranes, and cell walls. The trick in isolating pure plasmid DNA is to separate it from the rest of the cell components and from chromosomal DNA. The most common method used for separating plasmid DNA from chromosomal DNA is the alkaline lysis method developed by Birnboim and Doly (1979). They exploited the supercoiled nature of plasmid DNA to separate it from chromosomal DNA. Not only is plasmid DNA circular, it is also twisted by a bacterial topoisomerase. In the alkaline lysis method, both chromosomal and plasmid DNA are denatured (rendered single stranded) by NaOH, which disrupts hydrogen bonding. Denatured DNA can reanneal at neutral pH if it is not kept in alkali for too long. The two halves of the plasmid double-helix DNA remain intertwined during the incubation in alkali and they are in close proximity for reannealing. Chromosomal DNA is much larger, not intertwined, and often fragmented. Chromosomal DNA remains denatured and is precipitated by potassium acetate and sodium dodecyl sulfate (SDS), an ionic detergent. The precipitated chromosomal DNA is usually removed by filtration or centrifugation. RNA is also generally removed during the alkaline lysis step simply by adding RNase to the buffer. Plasmid DNA remaining in the supernatant can then be precipitated by ethanol. Steps 4 through 7 of the Qiagen protocol below represent the alkaline lysis portion of the purification protocol. The plasmid DNA is further purified over an anion-exchange column. Anion exchange chromatography works by taking advantage of the negatively charged nature of DNA. A matrix of positively charged molecules is immobilized in
13
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the column. When the cellular components are applied to the column, the negatively charged nucleic acids adsorb to the positively charged matrix. Proteins and other undesirable cellular debris are washed out of the column, and only the nucleic acids remain. The highly purified plasmid DNA can then be eluted from the column using a high-salt buffer. DNA quantification is accomplished by reading the absorbance of a known volume of sample at 260 nm. The absorption coefficient of pure DNA is 50 µg/ml. This means that one A260 unit of double-stranded DNA corresponds to 50 µg of DNA per ml. To assess the purity of a DNA sample, the ratio of the absorbance at 260 nm over the absorbance at 280 nm is calculated. A ratio of approximately 1.8 is ideal. A sample with a higher ratio may have RNA contamination, and a sample with a lower ratio may have protein contamination.
II. LABORATORY EXERCISES A. Alkaline Lysis and Anion Exchange Chromatography We will perform the plasmid purification using the QIAfilter Plasmid Maxi Kit. This and Promega’s Wizard DNA preparation kits are commonly used kits in molecular biology laboratories. The following protocol and Figure 2.1 have been adapted from the QIAGEN® Plasmid Purification Handbook (2003). 1. Two afternoons before your laboratory, your instructor picked a single colony of E. coli strain NovaBlue (or other K12 strain), containing either the pET-41a(+) or the pEGFP-N1 plasmid, from a freshly streaked LB/kanamycin plate. A starter culture of 2–5 ml LB medium containing kanamycin was inoculated and incubated for ~8 h at 37˚C with vigorous shaking (~300 rpm). Note: Use a snap cap tube or flask with a volume of at least 4 times the volume of the culture to provide adequate aeration. 2. The evening before your laboratory, your instructor diluted the starter cultures 1/500 to 1/1000 into 100 ml selective LB/kanamycin medium. Use a flask or vessel with a volume of at least 4 times the volume of the culture. The culture should reach a cell density of approximately 3–4 × 109 cells per ml, which typically corresponds to a pellet wet weight of approximately 3 g/liter medium. 3. Pour the E. coli cultures into centrifuge tubes and balance the tubes. 4. Harvest the bacterial cells by centrifugation at 6000 × g for 15 min at 4˚C. Remove all traces of supernatant by inverting the open centrifuge tube until all liquid has been drained. (Note: If you wish to stop the protocol and continue later, freeze the cell pellets at −20˚C. Don’t do this today, though.) 5. Resuspend the bacterial pellet in 10 ml Buffer P1. For efficient lysis it is important to use a vessel that is large enough to allow complete mixing of the lysis buffers. Ensure that RNase A has been added to Buffer P1. The bacteria should be resuspended completely by vortexing or pipetting up and down until no cell clumps remain. Buffer P1 is the resuspension buffer. 6. Add 10 ml Buffer P2, mix gently but thoroughly by inverting 4–6 times, and incubate at room temperature for 5 min. Do not vortex, as this will result in shearing of genomic DNA. The lysate should appear viscous. Do not allow the lysis reaction to proceed for more than 5 min. After use, the bottle containing Buffer P2 should be closed immediately to avoid acidification from CO2 in the air. Buffer P2 contains sodium hydroxide and a detergent and is used for cell lysis and denaturation of DNA. During the incubation prepare the QIAfilter Cartridge: Screw the cap onto the outlet nozzle of the QIAfilter Maxi Cartridge. Place the QIAfilter Cartridge in a convenient tube, such as a disposable 50 ml conical tube.
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15 7. Add 10 ml chilled Buffer P3 to the lysate, and mix immediately but gently by inverting 4–6 times. Proceed directly to step 8. Do not incubate the lysate on ice. Precipitation is enhanced by using chilled Buffer P3. After addition of Buffer P3, a fluffy white precipitate containing genomic DNA, proteins, cell debris, and SDS becomes visible. The buffers must be mixed completely. If the mixture still appears viscous and brownish, more mixing is required to completely neutralize the solution. It is important to transfer the lysate into the QIAfilter Cartridge immediately in order to prevent later disruption of the precipitate layer. Buffer P3 neutralizes the solution, causing plasmid DNA to reanneal, and acts to precipitate the single stranded chromosomal DNA. 8. Pour the lysate into the barrel of the QIAfilter Cartridge. Incubate at room temperature (15–25˚C) for 10 min. Do not insert the plunger! Important: This 10 min incubation at room temperature is essential for optimal performance of the QIAfilter Maxi Cartridge. Do not agitate the QIAfilter Cartridge during this time. A precipitate containing proteins, genomic DNA, and detergent will float and form a layer on top of the solution. This ensures convenient filtration without clogging. If, after the 10 min incubation, the precipitate has not floated to the top of the solution, carefully run a sterile pipet tip around the walls of the cartridge to dislodge it. 9. Equilibrate a QIAGEN-tip 500 by applying 10 ml Buffer QBT and allow the column to empty by gravity flow. Flow of buffer will begin automatically by reduction in surface tension due to the presence of detergent in the equilibration buffer. Allow the QIAGEN-tip to drain completely. QIAGEN-tips can be left unattended, since the flow of buffer will stop when the meniscus reaches the upper frit in the column. 10. Remove the cap from the QIAfilter Cartridge outlet nozzle. Gently insert the plunger into the QIAfilter Maxi Cartridge and filter the cell lysate into the previously equilibrated QIAGEN-tip. Filter until all of the lysate has passed through the QIAfilter Cartridge, but do not apply extreme force. Approximately 25 ml of the lysate are generally recovered after filtration. 11. Allow the cleared lysate to enter the resin by gravity flow. 12. Wash the QIAGEN-tip with 2 × 30 ml Buffer QC. Allow Buffer QC to move through the QIAGEN-tip by gravity flow. The first wash is sufficient to remove all contaminants in the majority of plasmid preparations. The second wash is especially necessary when large culture volumes or bacterial strains producing large amounts of carbohydrates are used. 13. Elute DNA with 15 ml Buffer QF. Collect the eluate in a 30-ml tube. Use of polycarbonate centrifuge tubes for collection is not recommended as polycarbonate is not resistant to the alcohol used in subsequent steps. 14. Precipitate DNA by adding 10.5 ml (0.7 volumes) room-temperature isopropanol to the eluted DNA. Mix and centrifuge immediately at 15,000 × g for 30 min at 4˚C. Carefully decant the supernatant. All solutions should be at room temperature in order to minimize salt precipitation, although centrifugation is carried out at 4˚C to prevent overheating of the sample. Isopropanol pellets have a glassy appearance and may be more difficult to see than the fluffy, salt-containing pellets that result from ethanol precipitation. Marking the outside of the tube before centrifugation allows the pellet to be more easily located. Isopropanol pellets are also more loosely attached to the side of the tube, and care should be taken when removing the supernatant. 15. Wash DNA pellet with 5 ml of room-temperature 70% ethanol and centrifuge at 15,000 × g for 10 min. Carefully decant the supernatant without disturbing the pellet. Alternatively, disposable conical-bottom centrifuge tubes can
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be used for centrifugation at 5000 × g for 60 min at 4˚C. The 70% ethanol removes precipitated salt and replaces isopropanol with the more volatile ethanol, which can be removed by evaporation. 16. Air-dry the pellet for 5–10 min, and dissolve the DNA in a suitable volume* of buffer (e.g., TE buffer, pH 8.0, or 10 mM Trisz.Cl, pH 8.5). (*For pEGFP-N1, use 300 ml, and for pET-41a(+), use 100 ml.) Dissolve the DNA pellet by rinsing the walls to recover all the DNA, especially if glass tubes have been used. Pipetting the DNA up and down to promote resuspension may cause shearing and should be avoided. Overdrying the pellet will make the DNA more difficult to dissolve. DNA dissolves best under alkaline conditions and is stable for long periods of time at pH 8. 17. Transfer to a microcentrifuge tube, mix well, and allow the DNA to resuspend for at least 10 minutes before proceeding to DNA quantification.
QIAfilter Plasmid Kit Pelleted bacteria
Alkaline lysate
Clear lysate by filtration
QIAfilter Midi QIAfilter Maxi
Bind DNA Wash Elute
Add Isopropanol
Collect DNA by centrifugation
Ultrapure plasmid DNA Fig. 2.1 QIAfilter Plasmid Maxi Kit Flowchart. Copyright 2004 Qiagen Corporation. Used with permission.
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Lab Session 2
B. DNA Quantification This method for DNA quantification uses the GeneQuant apparatus from Amersham Biosciences. It also uses a special cuvette that accepts a quantity as small as 70 µl. This cuvette allows for minimal sample waste. If this apparatus is not available, a standard uv/vis spectrophotometer can be used to assess the absorptions at 260 and 280 nm. 1. Turn on the GeneQuant (or spectrophotometer) 15 minutes before use. 2. In a microcentrifuge tube, add 2 µl DNA to 98 µl dH20. Mix well. This is a 50-fold dilution of your DNA. (Note: A 50-fold dilution of your DNA is not always appropriate. We are diluting 50 times because we assume that a very high concentration of DNA will be present due to the large volume of bacteria from which we started.) 3. Blank the GeneQuant (or spectrophotometer) using 70 µl of water in the quartz cuvette. Use nonpowdered gloves. Be especially careful not to get fingerprints on the clear side of the cuvette. If you think you have fingerprints on your cuvette, rinse and wipe well with a Kimwipe. 4. After the instrument has been blanked, carefully empty and rinse the cuvette. Note: Please be careful with the quartz cuvette. These cuvettes are shockingly expensive and are NOT disposable. 5. Read the absorbance of 70 µl of your diluted DNA at 260 nm and 280 nm. Record the readings in your notebook. 6. Empty and rinse cuvette. 7. Calculate the concentration and purity of your original sample. To determine the concentration of your DNA, use the equation (A260) × (50 µg/ml) × (dilution factor) = DNA (µg/ml) Remember, in this case, your dilution factor was 50. To determine the purity of your DNA: A260/A280 9. Record the DNA concentration and ratio in your notebook. If the A260/A280 ratio was significantly different from 1.8, see your instructor. 10. Label your original/undiluted tube of purified DNA with your station number, the name of the plasmid you purified, the concentration (with the units µg/ml), and the purity. The instructor will collect these tubes at the end of the period so that the DNA can be pooled and redistributed to you since each group will need both plasmids and you only purified one of the two today. Your instructor will run a portion of your DNA sample on a gel before your next laboratory period to check for degradation and to verify the concentration.
REFERENCES Birnboim, HC, and J Doly. 1979. A rapid alkaline extraction procedure for screening recombinant plasmid DNA. Nucl. Acids Res. (6), 1513–1523. QIAGEN® Plasmid Purification Handbook. August 2003. 2nd Ed. http://www1.qiagen.com/literature/ handbooks/PDF/PlasmidDNAPurification/PLS_Plasmid/102 5302_HB_PLS_082003WW.pdf.
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LAB SESSION
3 Preparation of Expression Vector DNA pET-41a(+), a GST Fusion Protein Vector oal: Today you will prepare the expression vector plasmid, pET-41a(+) (Novagen), to be able to accept the gene (egfp) you are going to clone in the following weeks. To accomplish this, you will digest pET41a simultaneously with two restriction endonucleases, Nco I and Not I. This will allow you to clone the egfp gene into the vector in a single orientation, ensuring correct translation of a GST::EGFP fusion protein. To orient you, here is a summary of the next series of experiments:
G
1. Cut vector pET-41a(+) (today). 2. Perform restriction digest of pEGFP-N1 and run preparative gel to isolate egfp fragment. 3. Ligate insert (egfp) to vector (pET-41a(+)). 4. Transform ligation products into competent cells. 5. Pick putative transformants and make replica plates. 6. Identify positive clones.
I. INTRODUCTION In general, cloning vectors are plasmids that are used primarily to propagate DNA. They replicate in E. coli to high copy numbers and contain a multiple cloning site (also called a polylinker) with restriction sites used for inserting a DNA fragment. A selectable marker gene, such as an antibiotic resistance gene, is included to ensure survival of the plasmid. A screenable marker, such as β-galactosidase, is also often included. An expression vector is a specialized type of cloning vector. Expression vectors are designed to transcribe the cloned gene and translate it into protein. They do have some features in common with the general cloning vectors that are used only for propagating DNA, such as the multiple cloning site and the selectable marker, but they tend to have a lower copy number within cells, and they rarely have a screenable marker. They also have some important additional features that allow them to express genes and make protein, including a promoter, ribosome binding site, ATG start codon, a multiple cloning site (polylinker) that allows inserts to be ligated in a predictable reading frame, and often (not always) a fusion tag to aid in purification steps.
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Expression Vectors The expression vector you will use for your project is pET-41a(+) (Figure 3.1). This expression vector utilizes a kanamycin resistance gene as a selectable marker and the glutathione-S-transferase gene (gst) as a fusion tag. The multiple cloning site is downstream (3’) of the gst gene, and there is no stop codon or termination signal following the gst gene. Therefore, when our gene of interest (egfp) is cloned into the multiple cloning site, it will be expressed as a fusion protein with gst, resulting in expression of the fusion protein, GST::EGFP. You will learn more about how creating this fusion protein will aid in the purification of the EGFP protein later in the semester. The expression of the gst gene, and consequently the fusion gene in your future construct is under the control of the T7 promoter and is inducible using isopropyl-ßD-thiogalactopyranoside (IPTG). In nature, the promoter is induced by lactose, and IPTG mimics lactose with regard to the induction properties, but is not cleaved by the E. coli enyme β-galactosidase. Inducibility is due to the fact that pET-41a(+) uses two components of the lac operon, the lac operator and the lacI gene, to regulate transcription. In this vector, the lac operator is located adjacent to the T7 promoter. lacI encodes a repressor and is constitutively expressed, so the repressor protein LacI is always present. LacI binds to the lac operator in the absence of inducer, and prohibits RNA polymerase from initiating transcription from the T7 promoter. When the inducer molecule IPTG is added, it interacts with LacI in such a way that LacI will no longer bind to the lac operator, and thus transcription by the T7 RNA polymerase proceeds. This process is called derepression of the promoter (Figure 3.2).
T7 terminator NotI NcoI (ATG) gst kan ATG T7 promoter lac operator pET-41a(+) 5933 bp
lac I
Fig. 3.1 Salient features of pET-41a(+)
Orientation To ensure that our gene of interest will be inserted in the proper orientation, we will employ the method of directional cloning, also called “forced” cloning (Figure 3.3). In forced cloning, the polylinker of the vector is digested with two different restriction endonucleases that leave incompatible cohesive ends, and the small “stuffer fragment” between the two restriction sites is removed. The insert (your gene of interest) is then cut out with the same two restriction endonucleases and ligated into the vector. Cloning in this manner, rather than cloning by cutting with only one restriction endonuclease has two advantages: 1. The incompatible cohesive ends will prevent vector from religating without incorporating the insert.
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Lab Session 3 A. Repressed state
B. Derepressed state due to inducer molecule
LacI gene of interest RNA polymerase
lac operator promoter
RNA polymerase
gene of interest lac operator promoter
LacI
LacI
lacI
lacI
IPTG
Fig. 3.2 Promoter repression by LacI and derepression by IPTG. A. The repressed state of the promoter. B. The derepressed state of the promoter due to the inducer molecule, IPTG. When cloning the gene of interest into an expression vector, it is critical for the gene both to be in the correct orientation and the proper reading frame with respect to the start of translation (the ATG start codon encoded by the vector).
2. The orientation of the insert is forced in a single direction: that is, the 5′ end of the gene can ligate with only one end of the cut vector. Because we know the sequence of both the vector and insert, we know that the fusion protein gene will be translated, or expressed, correctly. You will cut the vector with two restriction enzymes that have recognition sequences within the multiple cloning site: Nco I and Not I. Later, the egfp gene will be excised using the same two restriction enzymes (Nco I at the 5′ and Not I at the 3′ end) and it will be ligated into the expression vector.
Forced (Directional) Cloning 5'
3'
TTCAA
A
3'
5'
G
TTCGA 3'
5'
5'
G T CT AA
T
3' A
CT
AG
Fig. 3.3 An example of forced cloning using the EcoRI and HindIII restriction endonucleases. The insert can only be incorporated in one orientation.
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Part I. Manipulation of DNA
Remember that the Watson and Crick strands are anti-parallel. If Nco I was the only restriction enzyme used to both cut out the egfp insert and to cut the pET-41a(+) vector, the insert would have the ability to be incorporated into the vector in either orientation. In this case, only 50% of your transformants would contain the egfp gene in the proper orientation. Only transcription from DNA in the correct orientation will result in the correct mRNA and the correct amino acid sequence being produced. DNA can be transcribed only in a 5′ to 3′ direction, and the sequence on the bottom strand, 5′ to 3′, is different from the sequence on the top strand. See Appendix 3 to make sure you understand this.
Reading Frame The reading frame with respect to the translational start site must be maintained for correct expression. In pET-41a(+), the junction of foreign DNA with gst has to be in the proper reading frame to create the desired GST::foreign peptide fusion protein. Most expression vectors are designed in families of three members. Typically, all three expression vectors in the family are identical, except that the reading frames with respect to the multiple cloning sites differ. For example, for a given restriction site, the first vector may put an insert in the +1 reading frame with respect to gst, the second in the +2 reading frame, and the third in the +3 reading frame. Only one of the three vectors will maintain the correct reading frame for a given insert. The other two will result in the insert being in the wrong reading frame. pET-41a(+) and several other recently developed expression vectors have an additional feature of the multiple cloning site: an Nco I restriction site. The Nco I recognition sequence is useful in that it contains an ATG sequence, the start codon for most proteins. The complete Nco I recognition sequence is CCATGG. The Nco I recognition sequence in pET-41a(+) is located such that the ATG of the sequence is in-frame with the ATG start codon of the gst fusion tag. Therefore, if your gene of interest starts with an ATG that is part of an Nco I site, then the vector and the 5′ end of the insert (your gene of interest) can both be cut with Nco I, and the gene will automatically be in the correct reading frame for translation. Fortunately, the egfp gene that you will clone into pET-41a(+) does contain the Nco I recognition sequence at the ATG start of translation. Therefore, both the vector and the 5′ end of the insert may be cut with Nco I, and then ligated together with confidence that the insert will be in the correct reading frame. Illustrated below is a portion of the multicloning region of the pET-41 family of expression vectors. Note how the Nco I site is in the same reading frame for all of the vectors, but addition or deletion of a single base pair downstream of the Nco I site changes the amino acid sequence, while also setting up the downstream restriction sites in different reading frames. BamHI is highlighted as a reference site. The restriction site for Nco I is C/CATGG and the restriction site for BamHI is G/GATCC. pET-41a(+): ATG...gst...CC/C-ATG-GGA-TAT-CGG-G/GA-TCC-GAA-TTC Met Pro Met Gly Tyr Arg Gly Ser Glu Phe pET-41b(+): ATG...gst...CC/C-ATG-GAT-ATC-GGG/-GAT-CCG-AAT-TC Met Pro Met Asp Ile Gly Asp Pro Asn pET-41c(+): ATG...gst...CC/C-ATG-GCG-ATA-TCG-GG/G-ATC-CGA-ATT-C Met Pro Met Ala Ile Ser Gly Ile Arg Ile
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II. LABORATORY EXERCISES A. Restriction Digestion of Vector (pET41a) Restriction Enzyme Digestions Restriction enzyme activity is defined as the amount of enzyme (measured in units, U) that will cleave 1 µg of DNA (usually λDNA) to completion in 1 hr at the optimum temperature for the enzyme, usually 37˚C. Buffers are usually supplied with restriction enzymes at a 10× concentration. As a general rule, to set up a restriction enzyme digest, determine the amount of DNA to be cleaved, use a fivefold excess of enzyme, ensure that the volume of enzyme does not exceed 10–20% of the final volume, and add 10× buffer to a final concentration of 1×. For example, to digest 2 µg of DNA at a concentration of 0.5 µg/µl, the following reagents would be added, in order: DNA Water 10× buffer Enzyme (10 U/µl)
4 µl 13 µl 2 µl 1 µl
Total volume
20 µl
The reaction could also be set up with 3 µl of water and 1 µl of buffer for a total volume of 10 µl. Some enzymes will cleave at a second site under suboptimal conditions, producing what is referred to as “star activity.” For this reason, the enzyme should be diluted at least 1:10 and should be added last, to the diluted enzyme buffer. As a general rule, and depending on the number of restriction sites for a given enzyme and the size of the well in the gel, 100 to 200 ng is a good volume to load on an agarose gel for analysis. Thorough digestion of your vector is desired, therefore you may be encouraged to come either the afternoon before your lab or the morning of your lab to start your restriction digest. Each group will receive 2.5 µg of vector DNA. If the DNA concentration is 250 µg/ml, then the volume will be 10 µl. 1. Set your heat block to 65 degrees Celsius. There should be a thermometer in the heat block. This is for heat-inactivation of the enzyme after digestion. Your restriction digestion will be performed in a 37˚C incubator. 2. Immediately remove 2 µl (0.5 µg) and save in a tube labeled “pET41a uncut” and place it in your freezer box stored at −20˚C. 3. Digest the remaining DNA by adding the reagents listed below, being sure to add reagents in the order in which they are listed. Spin centrifuge tubes that contain small volumes of liquid for 5 seconds before removing aliquots—enzyme, DNA, buffer, etc. Make sure that buffers, which are stored at −20˚C are completely thawed before using. Note: This and other protocols use enzyme and buffer from New England Biolabs (Ipswich, MA). Other brands of restriction endonucleases may be used, but be sure to use the buffer suggested by that particular manufacturer at the concentration suggested by the manufacturer. Much of this information is available on the Web. ● ● ● ● ● ●
8 µl pET-41a(+) DNA 23.6 µl dH2O 4 µl 10X restriction buffer (NEB buffer 3) 0.4 µl BSA 2 µl Nco I (always add enzyme last) 2 µl Not I (always add enzyme last)
4. Mix and spin for 2–5 seconds to bring contents to bottom of tube. 5. Place tube in a microfuge rack in a 37˚C incubator for at least 2 hours. It is critical for the digestion to go to completion.
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B. Agarose Gel Electrophoresis Agarose gel electrophoresis works on the premise that all DNA is negatively charged. When DNA is added to the negatively charged pole (cathode) of an electrical field, it will move toward the positively charged pole (anode). Because of the pore size of the agarose matrix, large DNA molecules will be retarded in the matrix more than small molecules, so the small molecules will migrate more rapidly through the gel. Distinct bands ordered according to size can be visualized if the electric field is stopped before bands run off the end. You will run the digested DNA on an agarose gel to visualize whether the DNA cut to completion. Linear DNA migrates as a single band of a predictable size on an agarose gel. Circular plasmid DNA can appear as multiple bands and does not migrate at the same rate as linear DNA because of its secondary structure. In this exercise, you will compare the appearance of the linearized DNA from your reaction to an undigested sample. If your sample is not completely digested, there will be two or more bands instead of one. Make sure that you do not see the circular DNA in the digested sample to be sure that the digested DNA cut to completion. If you do have two or more bands, consult your instructor. You should pour your DNA gel during the restriction digestion. The teaching assistant or instructor will demonstrate how to pour a gel using your specific apparatus. You will each receive 1 ml of EtBr at a concentration of 1 mg/ml in a 1.5-ml microcentrifuge tube. Note: This tube should be saved and stored protected from light. CAUTION: Ethidium bromide (EtBr) is a carcinogen. Remember to wear gloves when working with ethidium bromide and to dispose of contaminated tips and gels in the containers specially marked for ethidium bromide solid waste disposal. 1. To prepare 1 X TBE buffer, mix 200 ml 5× TBE buffer with 800 ml water. 2. To prepare a 1% agarose gel, add 1 g of agarose to 100 ml of 1× TBE buffer. Microwave for 30 seconds with the cap on loosely. Swirl and repeat until agarose is completely in solution. Be sure to use rubber “hot hands” to handle the hot bottle, and never swirl the bottle close to your face in case the liquid boils over. CAUTION: Never leave the agarose solution unattended when using the microwave. The solution must be swirled occasionally during the heating process to prevent superheating of local areas. Always use “hot hands” or autoclave gloves when heating the agarose. 3. Apparati differ, but for mini-gels using the Gibco BRL Horizon 58, 25–30 ml of agarose is sufficient for pouring the gel. Allow the agarose to cool slightly before pouring the gel, because steaming hot agarose can warp the gel apparatus. EtBr must be added to both the agarose before it solidifies and to the TBE running buffer. Pour 30 ml of your melted agarose into a disposable 50-ml conical test tube. Then, ● ●
Add 15 µl ethidium bromide to the 30 ml agarose Add 16 µl ethidium bromide to 160 ml TBE
Allow the remaining agarose to solidify in the bottle with the cap tightly closed. This agarose will be re-melted in the microwave for pouring gels during the next several exercises. Before running the gel, loading buffer must be added to your sample. Loading buffer is also called “sample buffer,” “loading dye,” and “blue juice.” It contains glycerol to make the solution heavy so it will sink to the bottom of the well and bromophenol blue for visualization. If your sample is small, you may also add water to increase the volume to make loading easier. 4. To prepare your digested DNA sample for loading on the gel, mix together in a separate tube:
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2 µl of the digested pET-41a(+) 7 µl sterile distilled water 1 µl 10× loading dye
Mix, then centrifuge for 2 seconds. Leave the rest of the restriction digest at 37˚C while running the gel. This will be the DNA you will use in following weeks for cloning! Do not add loading dye to this sample: if you do, it can’t be used for cloning. 5. To prepare uncut DNA for loading on the gel, mix together in a separate tube: ● ● ●
0.5 µl uncut DNA (save the rest in freezer box for a later laboratory) 8.5 µl sterile distilled water 1 µl 10× loading dye
Mix, then centrifuge for 2 seconds. 6. Load your gel as follows: ● ● ● ● ●
Lane 1: Empty Lane 2: 5 µl Invitrogen 1 kb ladder (premixed with loading dye) Lane 3: Your digested DNA sample described above Lane 4: Empty Lane 5: Uncut DNA described above
The Invitrogen 1 kb ladder is a molecular weight marker for estimating DNA size. The visible bands are approximately 12, 11, 10, 9, 8, 7, 6, 5, 4, 3, 2, 1.6, 1, 0.5, 0.4, 0.35, and 0.3 kilobases in size (Figure 3.4).
1Kb DNA Ladder 1% agarose gel stained with ethidium bromide bp 11,198 9,162 7,126 -
- 12,216 - 10,180 - 8,144 - 6,108
5,090 4,072 3,054 -
2,036 1,636 -
1,018 -
506, 517* 396* 344* 298* 220* 201* 154* 134* 75*
* Hinf I fragments of the vector
0.5 µg/lane Fig. 3.4 Invitrogen 1 kb ladder. Copyright 2004 Invitrogen Corporation. Used with permission. Note that the 1.6 kb band is somewhat brighter than the adjacent bands. This is done to provide an easy marker for mw.
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7. Once you have loaded the samples, make sure that 1× TBE buffer covers the gel. Close the apparatus and connect the leads to the voltage pack, as demonstrated by your instructor. Run your DNA gel at approximately 85 volts, or at an appropriate voltage for your apparatus (10V/cm). Once the bromophenol blue dyefront runs to the bottom half of your gel, stop the current and place the gel on a UV transilluminator. Wear UV protective goggles whenever direct exposure to UV is a possibility. Photograph the gel and examine the photograph. Your instructor will help you determine whether your plasmid cut to completion. There should be a single DNA band in your cut DNA sample lane, and it should run at a slightly different size than uncut DNA.
C. Cleaning DNA Using a Spin Column The vector you linearized in this lab will be used in a later laboratory for ligation with the insert (egfp) DNA. Ligations are very sensitive to salt concentrations, so it is important to remove the salts used in the restriction digestion buffer. It is also necessary to inactivate the restriction enzymes (so they cannot “undo” the ligation during or after ligation), and to remove the stuffer fragment (the small piece of DNA between the 2 restriction sites). Heat Kill the Restriction Enzymes: 1. To “kill” the restriction enzyme activity, fill a heat block with water, and heat to 65˚C. Heat sample for 15 minutes, watching to be sure the top doesn’t pop open, allowing sample to evaporate or become contaminated. Note that not all restriction enzymes can be inactivated by this treatment. This kind of information can be found online at technical support sites for most suppliers. 2. Place sample on ice for 2 minutes to allow steam to condense, then centrifuge sample into the bottom of the tube for 5 seconds. Make sure to balance the microcentrifuge with an empty microcentrifuge tube. Remove Salts and Stuffer Fragment We will use the Qiagen QIAquick PCR Purification Kit Protocol to remove the salts, heat-inactivated enzymes, and stuffer fragment from our linearized vector. This kit was designed to remove enzyme, salts, and PCR primers from PCR reactions, but also serves our purpose because the stuffer fragment is similar in size to PCR primers. A large percentage of the stuffer fragment is removed because its small size inhibits its ability to bind in the spin column, and so most of it is lost in the wash steps. The spin column works on the principle of anion exchange chromatography, just like the column used during the large-scale plasmid prep last week. Alternatively, one could achieve the same goal by performing an ethanol or isopropanol precipitation, but the QIAquick protocol is prefered because it is more rapid and more efficient (less sample is lost) than an alcohol precipitation. The protocol and diagram (Figure 3.5) below are modified from the manufacturer’s handbook (QIAquick Spin Handbook 03/2001). Note: All centrifugation steps are performed at ~ 13,000 rpm in a conventional microcentrifuge. Balance your tube(s) with that from another lab group. 1. Add 190 µl Buffer PB to your sample and mix. 2. Place a QIAquick spin column in the 2-ml collection tube. 3. To bind DNA, apply the sample to the QIAquick column and centrifuge 30 sec. 4. Discard the flow-through. Place the QIAquick column back in same collection tube. 5. To wash, add 0.75 ml Buffer PE to the QIAquick column, and centrifuge 30 sec.
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6. Discard flow-through and place the QIAquick column back in same collection tube. Centrifuge an additional 1 minute. 7. Place the QIAquick column in a clean, sterile 1.5-ml microcentrifuge tube. Cut the lid off the 1.5-ml tube with scissors so that it will fit in the microcentrifuge. 8. To elute the DNA, add 50 µl Buffer EB to the center of the QIAquick membrane and centrifuge for 1 min. 9. Transfer your purified DNA to a fresh microcentrifuge tube labeled pET41a/Nco I/Not I and save in your freezer box.
QIAquick spin in microcentrifuges restriction digestion, PCR or other enzymatic reaction or solubilized gel slice
bind
wash
elute
Pure DNA fragments
Fig. 3.5 QIAquick PCR Purification Kit Flowchart. Copyright 2004 Qiagen Corporation. Used with permission.
REFERENCES QIAGEN® QIAquick Spin Handbook. July 2002. http://www1.qiagen.com/literature/handbooks/ PDF/DNACleanupAndConcentration/QQ_Spin/1 021422_HBQQSpin_072002WW.pdf.
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4 Preparation of Insert DNA (egfp) oal: Today you will excise the egfp gene from the plasmid pEGFP-N1 using the Nco I and Not I restriction enzymes. You will then gel purify the DNA fragment containing the egfp gene from the remainder of the plasmid. This will produce the pure egfp gene with Nco I sticky ends at the 5′ and Not I sticky ends at the 3′ end to use as insert for the ligations next week. Recall that the two enzymes used to cut out the egfp gene are the same two enzymes used last week to linearize the expression vector, pET-41a(+). This will enable a forced cloning where the insert can only be ligated to the vector in the correct orientation.
G I. INTRODUCTION
The gene we will be cloning and expressing is egfp (the gene encoding the enhanced green fluorescent protein). The green fluorescent protein (gfp) is a naturally occurring protein found in a species of fluorescent jellyfish called Aequoria victoria. The difference between the fluorescence of the green fluorescent protein (GFP) and the enhanced green fluorescent protein (EGFP) is that EGFP emits 35 × the fluorescence (at 510 nm) of GFP when excited with ultraviolet or blue light (395 nm, with a minor peak at 470 nm): it is much brighter. The increased fluorescence was achieved by making mutations in the nucleic acid sequence that resulted in a small change in the amino acid composition within the chromophore region of the protein: Ser65 -> Thr, and Phe64 -> Leu (Yang and Kain, 1996). In order to isolate the egfp gene from pEGFP-N1, we will perform a restriction digestion, followed by fragment separation and isolation on an agarose gel, followed by elution and purification of the DNA using glass milk. The egfp gene is flanked by an NcoI restriction site at the 5′ end, and a NotI restriction site at the 3′ end. The situation is complicated, however, by the fact that NcoI cuts the plasmid at 3 additional sites. The fragments arising from an NcoI/NotI double digest with their sizes and their cohesive ends are listed below, with the egfp gene shown in bold. Fragment Size 1904 1085 724 703 317
Cohesive Ends NcoI/NcoI NotI/NcoI NcoI/NotI NcoI/NcoI NcoI/NcoI
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The complication is that the desired 724 bp fragment cannot be easily separated from the 703 bp fragment by gel electrophoresis. Since the 703 bp fragment does not have a NotI cohesive end, however, it cannot be incorporated into the insert instead of the 724 bp fragment, so we will not worry about the co-purification of this fragment. In rare cases, it is possible that the 703 bp fragment could insert into the vector if it were ligated in tandem with the 724 bp fragment. This could be detected by various methods, including PCR, restriction mapping, and southern blot.
II. LABORATORY EXERCISES A. Restriction Digestion of pEGFP-N1 Because of time constraints, your instructor will set up the following restriction digest for each group, or your instructor may ask you to come either the day before lab, or several hours before lab to set up the digest. ● 30 µl pEGFP-N1 DNA (1 µg/µl concentration, therefore 30 µg total) ● 49 µl dH O 2 ● 10 µl restriction enzyme buffer ● 1 µl BSA ● 5 µl Nco I5 µl Not I Incubate at 37˚C overnight.
B. Isolation of egfp DNA from Agarose When you arrive in lab, set the heat block to 55˚C and fill with it water. Gel Separation of DNA Fragments We will not use ethidium bromide in this gel or in the buffer. Instead, we will incorporate Crystal Violet dye into the gel. The advantage of using Crystal Violet is that the DNA can be visualized without exposing the gel to UV light (as is necessary with ethidium bromide). This is valuable when isolating DNA from a gel because UV irradiation breaks down DNA, which is obviously a disadvantage when attempting to isolate a piece of DNA in order to clone it. The main reason Crystal Violet is not used routinely when viewing DNA gels is that it is not as sensitive as ethidium bromide, and only very high concentrations of DNA can be visualized. For this reason also, we will not run the 1 kb ladder on the prep gel. Although it would be useful, a normal concentration of 1 kb ladder cannot be visualized by Crystal Violet staining. 1. Prepare 1% agarose in TBE (swirl at frequent intervals during microwaving) 2. Obtain an aliquot of 1 mg/ml Crystal Violet and add 300 µl to 30 ml of the molten agarose. Swirl to mix thoroughly without causing bubbles. 3. If you do not have a special gel casting comb meant for preparative gels, use labeling tape to tape the teeth of your comb together to create one very long well in the gel. Be sure the tape is on tight and doesn’t hang off the end of the comb. 4. Pour the gel as usual, being sure the agarose is not so hot that it melts the glue off of the tape. 5. Once the gel is thoroughly solidified, remove the comb carefully. Prep combs have more of a tendency to rip the gel than regular combs. 6. Before loading your sample into the well, mix 30 µl of water with 3 µl of loading dye and load it into the prep well. Check to ensure that the water/dye sample does not leak through the bottom of the gel. If it does, then recast the gel and try again. If the test solution is retained in the well, proceed with loading your sample. There is no need to remove the test solution.
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7. Add 10 µl of loading dye to the 100 µl restriction digestion performed in part A. 8. Load this entire 110-µl sample on the gel, trying to distribute the sample evenly across the long prep well. 9. Run the gel until the smallest molecular weight band is approximately halfway through the gel, but do not run past that point. This gel should not be run at high voltage, because bands will smear. For the Horizon 58 gel box, run at 85 V. Do not run your gel more than half-way. Crystal Violet is a positively charged molecule, so while your negatively charged DNA travels down the gel, the Crystal Violet will travel “up” the gel. You may notice a region at the bottom of the gel that loses its purple color, and this region will grow in size the longer the gel is run. If you run the gel for too long, and a DNA band crosses the “de-stained” region of the gel, it will be very difficult to visualize the band. GENECLEAN® Protocol (modified from BIO 101® Systems handbook from Qiogene; see Appendix 2 for purchasing information) The following protocol takes advantage of DNA’s propensity to bind to glass. The glass milk consists of tiny glass beads in a buffered solution. Because of the high surface area to volume ratio of the glass milk, it is able to bind DNA very efficiently. Once DNA is bound, the agarose and TBE buffer salts can be washed away, and pure DNA can be eluted with a low-salt solution. 1. Excise 724 bp egfp DNA band from Crystal Violet-stained agarose gel with a razor blade. Lay the gel on a sheet of clean plastic to cut. Visualization may be increased by placing a white sheet of paper underneath the plastic or by placing it on a white light box. The 724 bp band is the third one from the top of the gel. Since the molecular weight is about .7 kb, it would run between the .5 and 1 kb molecular weight markers on the 1 kb ladder. Trim away any excess agarose to make the gel slice as small as possible without cutting away any of the DNA. Your gel should look similar to that in Figure 4.1. 2. Weigh the excised band and place in a microfuge tube. Weigh the gel slice (tare balance using an empty tube) to determine the approximate volume of the gel slice (0.1 g equals approximately 100 µl). If the gel slice weighs less than 0.25 g, use a 1.5-ml microcentrifuge tube; if the gel slice weighs more than 0.25 g, use multiple tubes (the volume needed, in ml, will be over 5 times the weight of the gel slice). It is not necessary to crush the gel, but large pieces can be sliced into roughly 2-mm cubes to facilitate dissolving the gel during the next step. If you have trouble understanding how to calculate your gel volume, ask your TA or instructor before adding NaI or TBE modifying buffer. 3. Add 4.5 volumes of NaI solution and 1/2 volume of TBE modifying buffer to the gel slice(s). 4. Melt agarose. Place the tube in a 55˚C water bath incubator/heat block. After a minute, vortex the contents of the tube and return it to the water bath. Continue the incubation until all of the agarose has dissolved (approx. 5 minutes). 5. Add glass milk. Resuspend glass milk by pipetting up and down. Add 5 µl glass milk per tube (if 2 tubes, 10 µl total) 6. Bind DNA to glass milk. After adding glass milk to the solution, mix (manually—do NOT vortex) and incubate at room temperature for 5 minutes to allow binding of the DNA to the silica matrix. Mix every 1 to 2 minutes to ensure that the glass milk stays suspended. If the volume of the binding reaction per tube is greater than 1 ml, allow at least 15 minutes while frequently mixing by hand, mechanical rocker, or rotation wheel to keep the silica particles in suspension. Vortexing the glass milk/DNA mixture will shear your DNA.
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0.7 kb
Fig. 4.1 Crystal violet-stained agarose gel of Nco I/Not I-digested pEGFP-N1
7. Pellet the silica matrix with the bound DNA. Microcentrifuge for approximately 5 seconds at full speed. Remove and save the supernatant in case all of the DNA did not bind to the glass milk. 8. Wash pellet 3 times with prepared NEW Wash (ethanol wash). Add 200 µl NEW Wash to the pellet (in each tube). The consistency of the pellet is different in NEW Wash (which is 50% ethanol) than in aqueous solutions and is somewhat resistant to resuspension. Resuspend the pellet in the wash by pipetting back and forth while digging into the pellet with the pipet tip. (Normally, pipetting up and down is not recommended for resuspending DNA because it can shear (break) DNA strands and if the DNA was greater than 15 kb, the glass milk procedure would not be advisable. In our case, the DNA bound to the glass beads is less likely to be sheared, and it is the only way to get the beads into suspension.) After it is resuspended, spin for 5 seconds in the centrifuge and discard the supernatant. Repeat the wash procedure two more times. After the supernatant from the third wash has been removed, spin the tube again for a few seconds and remove the last bit of liquid (with a small bore pipet tip) to avoid diluting the eluate with NEW Wash. Leave the cap open for 5–10 minutes at room temperature to dry the pellet. 9. Elute DNA from glass milk. Resuspend the washed, white pellet with 8 µl TE (per tube, if more than one tube). Centrifuge for about 30 seconds to make a solid pellet. Carefully remove the supernatant containing the eluted DNA and place in a new tube. This supernatant contains your egfp insert DNA. Label the tube with the supernatant “egfp insert.”
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Analytical Gel to Observe Purified egfp Fragment This gel serves two purposes: 1) to ensure that the egfp DNA was purified and that no other DNA fragments remain and 2) to estimate concentrations of both the vector and the insert DNA. A DNA mass ladder in which each band has a different amount of DNA is used to estimate the concentrations of your vector and insert (Fig 4.2). The DNA mass ladder that you are using contains an equimolar mixture of DNA fragments of 2000, 1200, 800, 400, 200, and 100 base pairs. Electrophoresis of 4 µl of the mixture results in DNA bands containing 200, 120, 80, 40, 20, and 10 ng, respectively (Invitrogen, 2004). You will compare the intensity of your vector and insert DNA to the intensity of the various bands in the ladder. Say, for example, you load 4 µl of the ladder on your gel, and you load 2 µl of a DNA sample of unknown concentration. If the band of the unknown sample is of the same intensity as the 400 base pair band in the ladder. The 400 bp fragment in the marker corresponds to 40 ng, so that would mean that there is 40 ng of the unknown DNA in the gel. Since you loaded 2 µl of that DNA, your original concentration of unknown DNA is 40 ng in 2 µl: in other words, 20 ng/µl. Be sure that all DNA samples go into and stay in the wells. If the sample or DNA mass ladder floats out of the well, then the estimation will not be accurate. The GeneQuant (or any spectrophotometric method) cannot be used for quantification of the insert because trace amounts of glass milk left behind in the sample will scatter light and interfere with the reading. Low DNA Mass Ladder 2% agarose gel stained with ethidium bromide bp
ng
2,000 1,200 -
- 200 - 120
800 -
- 80
400 -
- 40
200 -
- 20
100 -
- 10
4 µl/lane
Fig. 4.2 The Invitrogen low DNA mass ladder contains an equimolar mixture of DNA fragments of 2000, 1200, 800, 400, 200, and 100 base pairs. Electrophoresis of 4 µl of the mixture results in DNA bands containing 200, 120, 80, 40, 20, and 10 ng, respectively. Copyright 2004 Invitrogen Corporation. Used with permission.
For an alternative, less expensive way to estimate DNA concentration on a gel, λ HindIII fragments could be used (see manufacturer’s details: New England Biolabs catalog # N3012S). The concept is the same. 1. Prepare and run a 1% agarose gel with ethidium bromide as described in section 3B, but with the following samples: ● ●
Lane 1: 7 µl 1 kb ladder (pre-mixed with loading dye) Lane 2: Low DNA mass ladder (4 µl + 1 µl loading dye, then load all 5 µl)
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●
Lane 3: Digested vector (pET41a) from lab session 3 (mix 2 µl sample, 7 µl water, and 1 µl loading dye then load the total 10 µl mixture in the well) Lane 4: Purified egfp insert (mix 2 µl sample, 7 µl water and 1 µl loading dye then load the total 10 µl mixture in the well)
BE SURE TO SAVE YOUR VECTOR AND PURIFIED egfp INSERT IN YOUR FREEZER BOX AFTER REMOVING THE AMOUNT TO BE LOADED ON THE GEL. 2. Photograph the gel and estimate the intensities of vector and insert bands based on the DNA mass ladder. Since you loaded 2 µl of each of your samples of unknown concentration, once you determine the amount of DNA on the gel, you must divide by 2 to obtain the concentration in ng/µl units. 3. Calculations for next week: (YOU MUST HAVE THIS READY FOR THE LAB PERIOD NEXT WEEK) Calculate # of µl of your digested pET41a to equal 50 ng. Calculate # of µl of your purified egfp insert to equal 7 ng, and to equal 21 ng.
REFERENCES Invitrogen Low DNA Mass Ladder. Copyright 2004 Invitrogen Corporation. Used with permission. http://www.invitrogen.com/content/sfs/manuals/10068013.pdf Yang, T, L Cheng, and SR Kain. 1996. Optimized codon usage and chromophore mutations provide enhanced sensitivity with the green fluorescent protein. Nucl. Acids Res. 24 (22), 4592–4594.
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5 Preparation of Transformation-Competent Cells and Control Transformation oal: Today you will prepare Escherichia coli to become competent for DNA uptake. You will test the transformation efficiency of your cells using a standard preparation of supercoiled DNA.
G I. INTRODUCTION
Why and how bacterial cells can be made to be competent for DNA uptake by treatment with divalent cations is not understood. Fortunately, they can be made competent and they can be stored in this state indefinitely at −80˚C. Many parts of the procedure for making competent cells are critical for obtaining cells with a high transformation efficiency. The cells must be harvested in log phase, and once chilled they must not warm to room temperature or they will lose competence. For optimal transformation efficiencies, the cells should be aliquoted in a cold room. Because the expression vector we are using, pET-41a(+), has a T7 promoter that drives transcription of the gene of interest, the E. coli host strain must be a λDE3 lysogen so that it will have a T7 RNA polymerase to bind at the promoter for expression. A suitable strain is BL21(DE3).
II. LABORATORY EXERCISES A. Preparation of Chemically Competent Cells by Calcium Chloride Treatment Remember that you are not using antibiotics in this procedure. Use sterile technique and make sure the centrifuge bottles are sterile. The instructor will start an overnight culture of E. coli by using an inoculating loop to scrape cells from a single colony into 2 ml of LB in a polypropylene snapcap tube the evening before your laboratory. (Note: Antibiotics are not added.) They will be incubated at 37˚C overnight in a shaking incubator. 1. Start a 100-ml cell culture of E. coli (BL21 (DE3) or another suitable strain) with a fresh overnight culture as inoculum. Each group will inoculate one 1liter flask containing 100 ml of Luria-Bertani (LB) broth with 0.1 ml of overnight culture (1:1000 dilution).
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2. Incubate the flask at 37˚C with shaking. Grow the bacteria to log phase. An OD600 value between 0.3 and 0.5 may be used. It will take 2.5–4 hr for the culture to grow to the proper stage for making competent cells. 3. Using sterile technique and a sterile 5-ml pipette, withdraw samples from the cultures and determine the OD600 using a spectrophotometer (Spectronic 20 or equivalent). Once cultures appear cloudy, they are in log phase growth and their OD will increase rapidly. 4. When the proper optical density is reached, pour the culture into two sterile Oakridge tubes. Balance the tubes and then place on ice for 15 min. From now on, it is important that the cells never warm to room temperature. 5. Harvest the cells by centrifugation at 6000 rpm for 10 min at 4˚C. Make sure the rotor is prechilled. 6. Pour off the supernatant and, keeping the cells on ice, remove excess LB medium with a Pasteur pipette or a 1-ml micropipette. Resuspend the cells in a 1/100 volume of ice-cold sterile transformation and storage solution (TSS) (Chung et al., 1989). Resuspend gently by pipetting up and down. Transfer the contents of one tube to the other to consolidate your cells. Be very gentle here; the cells are now fragile. TSS contains 50 mM MgCl2; other procedures use 50 mM CaCl2. The dimethylsulfoxide (DMSO) in TSS is necessary if you intend to freeze some cells. Because there is little loss in transformation efficiency, it is always useful to freeze extra cells. 7. Pipette 100-µl aliquots quickly into cold microcentrifuge tubes. These cells can be used immediately or frozen at −80˚C. You should keep two tubes on ice for today’s test transformation and freeze the rest. You will need 6 tubes of frozen competent cells for next week. 8. To make frozen competent cells, aliquot 100 µl cells into cold microcentrifuge tubes (you will need at least eight tubes). Take your ice bucket to a dry/ice ethanol bath near a −80˚C freezer for storage of the cells. Using a precooled microcentrifuge tube holder or one layer of cheesecloth, submerge the tubes. Retrieve the tubes into a labeled cardboard or styrofoam box and immediately place at −80˚C. The transformation efficiency is measured by determining the number of colony-forming units (cfu)/µg DNA. Only supercoiled DNA should be used (linear DNA does not transform efficiently). In the following section you will use 10 ng of uncut pET-41a(+) DNA to measure the transformation efficiency of your cells.
B. Transformation Control 1. Label the tops of the two non-frozen tubes of competent cells (from Section II,A above) with your group number and a designation for the following treatments: a. TE, 10 µl b. pET-41a(+), 10 ng Remember to keep the cells on ice at all times. 2. Add the appropriate DNA to each tube, finger flick gently, and leave on ice for 30 min. If you are using cells stored at −80˚C, wait until the cells have thawed on ice before adding the DNA. 3. Add 900 µl of liquid medium (either TSS, LB, or SOC can be used), gently invert the tubes to mix, and incubate at 37˚C, 225 rpm for 60 min. This time is necessary to allow the antibiotic resistance gene to be expressed.
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5. Vortex the cells and plate 100 µl on selective medium (LB/kan). Be sure to label the backs of the plates with your lab day, station number, and the name of the sample. The remainder can be stored at 4˚C and plated at a later date if desired. (If you freeze the cells, make sure they have enough DMSO.) Always label the backs of Petri dishes rather than the lids to avoid confusion. 6. Incubate the plates at 37˚C overnight. 7. If the recovery of transformants is low, the remaining cells in 900 µl of cells can be concentrated by centrifugation and plated. Efficiencies are usually 1–4× 106 cfu/µg supercoiled pET-41a(+).
REFERENCES Chung, CT, SL Nienela, and RH Miller. 1989. One-step preparation of competent Escherichia coli: Transformation and storage of bacterial cells in the same solution. Proc. Nat. Acad. Sci. U.S.A. 86, 2172–2175.
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6 DNA Ligation and Transformation of Escherichia coli
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oal: The goal of this week’s laboratory is to ligate the egfp gene (our insert) into the linearized pET-41a(+) expression vector. We will then transform the ligation mixture into E. coli host cells.
I. INTRODUCTION DNA ligase catalyzes covalent bond formation between the 3′OH and 5′PO4 on DNA (phosphodiester bond). It requires two ends of double-stranded DNA and a buffer that includes ATP. Ligation kinetics are complex and it is worthwhile understanding some of the parameters that affect both the frequency and products of ligation. Bacteriophage T4 DNA ligase is the preferred enzyme for DNA ligation because it can efficiently ligate blunt-ended DNA as well as DNA ends with “overhangs,” short 5′ or 3′ extensions of single-stranded DNA created by restriction enzyme digestion. DNA ligase activity is measured in Weiss units: 1 Weiss unit is defined as the amount of enzyme that catalyzes the exchange of 1 nmol of 32P from pyrophosphate into ATP in 20 min at 37˚C. An amount of T4 DNA ligase equal to 0.015 Weiss unit will ligate 50% of the HindIII fragments of 5 µg bacteriophage lambda DNA, in 30 min at 16˚C. The optimal ligation temperature for a given reaction has been shown empirically to vary. DNA with compatible ends is the most amenable to ligation; 1–4 hr at room temperature or overnight at 4–15˚C is sufficient to catalyze the ligation of most ends. Blunt-ended DNA is ligated at room temperature or at 37˚C, the optimal temperature for the enzyme. Different preparations of ligase and ligase buffer call for differing ligation times at varying temperatures. Always be sure to check with manufacturers’ instructions. Ligation reaction temperatures are a balance between the activity of enzyme and annealing temperature of the compatible ends. The formula for calculating the melting temperature (TM) for short strands of DNA is TM = (2)(number of A + T pairs) + (4)(number of G + C pairs). As an exercise, calculate the annealing temperature for a 4-bp HindIII single-stranded DNA overhang (GATC). When designing ligation reactions, there are several practical rules of thumb that are useful to follow: ●
Ligation reactions are generally set up in small volumes of 10–20 µl so that compatible ends will not be too dilute in solution.
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For average sized vectors (2–6 kilobases), use approximately 50 ng of vector DNA per reaction. Dugaiczyk et al. (1975) empirically derived a formula for determining how much vector is ideal in a ligation reaction.
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The ideal amount of insert to use can only be determined empirically, but generally, try adding insert at a 1:1, 1:3, and 1:10 vector to insert molar ratio. F or example, if you are using a 3 kb vector and a 3 kb insert: ● ● ●
1:1 = 50 ng vector + 50 ng insert 1:3 = 50 ng vector + 150 ng insert 1:10 = 50 ng vector + 500ng insert ●
If you are using a 3 kb vector and a 1 kb insert:
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1:1 = 50 ng vector + 16.7 ng insert 1:3 = 50 ng vector + 50 ng insert 1:10 = 50 ng vector + 166.7 ng insert
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Ligase buffer includes the necessary salts and ATP, and is usually supplied as a 10× solution by the ligase manufacturer. Be sure to include it in your ligation mix or the ligase will not be active. 1 µl ligase per reaction from most commercially available sources is sufficient; however, a higher concentration of enzyme may be useful for blunt-end ligations.
For today’s experiment, we will attempt to ligate the expression vector with the egfp insert using 2 different ratios: a 1:1 and a 1:3 vector to insert ratio. We will also perform several control ligations and mock ligations (Figure 6.1). There are two pieces of information that are critical to understanding these controls. 1. DNA strands do not spontaneously ligate, even if the strands have complimentary cohesive ends. An enzyme that catalyzes phosphodiester bond formation (such as DNA ligase) must be present for ligation to occur. 2. Only circular DNA is transformable into competent E. coli cells. Linear DNA can get into the cells, but it cannot replicate once taken up. The first column of Figure 6.1 shows the type of DNA to be added to the control: circular or linearized. The second column indicates whether DNA ligase is added to the control. The third column shows whether that DNA should transform or not (circular DNA can transform, linear cannot). If the plasmid transforms, then transformed cells should grow on medium containing kanamycin since the vector contains the kanamycin resistance gene. The last column shows some explanations for unexpected results (where colonies grew when they shouldn’t have, or vice versa). Keep in mind that restriction digestions and ligation reactions rarely go to completion. It is normal to see a few background colonies on plates where you should not see transformants, even if the gel of ligation products appears as it should. These colonies will not contain insert. These colonies are present because the sensitivity of the transformation is much higher than the sensitivity of visualizing DNA molecules on an agarose gel. Also keep in mind that any DNA fragments with compatible cohesive ends may be ligated together. This means that you might see transformants that have one vector plus one insert, transformants with one vector and multiple inserts, vector with incorrect inserts, and multiple vector combinations. Although they do occur during the ligation reaction, it would be unlikely to recover insert-only, or concatamerized insert with no vector from a transformant, because such DNA molecules would be unlikely to contain an origin of replication, and therefore could not replicate in vivo. Question: You perform a ligation with NcoI/NotI-digested, pET-41a(+) vector and the egfp insert from pEGFP-N1 (as in lab). You then transform the ligation mix into E. coli and plate on LB medium containing kanamycin. Consider the following unexpected outcomes, and suggest controls: 1. Nothing grows. What controls might you design to determine whether the E. coli cells are viable (alive) versus whether the E. coli is competent? What would the results of those controls be?
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DNA treatment uncut pET41a R kan
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with ligase 1. investigate controls 2. bad ligase? 3. bad competent cells?
Fig. 6.1 DNA ligation and transformation controls
2. You see lots of growth, but when you isolate plasmid from numerous E. coli colonies, all you find is pET-41a(+) with no insert. Suggest the two most likely ways these colonies arose (aside from contamination). Also suggest controls to distinguish each possibility. What would the results of those controls look like?
II. LABORATORY EXERCISES A. Ligations and Ligation Controls Set up 6 ligations (or mock ligations) in microcentrifuge tubes. Label clearly with lab day, station number, and sample number. Ligation 1. 1:1 molar ratio pET-41a(+) : egfp insert 50 ng pET-41a(+) NcoI/NotI 7 ng egfp insert sterile dH2O 10× ligase buffer Ligase
µl µl µl 2 µl 1 µl 20 µl total
Ligation 2. 1:3 molar ratio pET-41a(+) : egfp insert 50 ng pET41a(+) NcoI/NotI 21 ng egfp insert sterile dH2O 10× ligase buffer ligase
µl µl µl 2 µl 1 µl 20 µl total
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Ligation 3. Linear pET-41a(+) no ligase control to control for the presense of uncut vector 50 ng pET-41a(+) NcoI/Not I sterile dH2O
µl µl 20 µl total
Ligation 4. Linear pET-41a(+) plus ligase control to control for presence of stuffer fragment and recircularization of vector 50 ng pET-41a(+) NcoI/Not I sterile dH2O 10× ligase buffer ligase
µl µl 2 µl 1 µl 20 µl total
Ligation 5. Circular (uncut) pET-41a(+) control to control for viability/competency of cells 50 ng pET-41a(+) uncut sterile dH2O
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Ligation 6. Sterile TE buffer control to control for contamination sterile TE buffer
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Mix gently, and bring contents to the bottom of the tube either by a 5-second centrifugation or tapping the contents to the bottom of the tube. Incubate for 10 minutes at room temperature. Note: This protocol uses NEB (New England Biolabs) T4 DNA ligase, catalog number M0202L. If using a different brand of T4 DNA ligase, follow manufacturer’s instructions. The incubation time may be longer. Proceed to transformation.
B. Divalent Cation-Mediated Transformation Competent cells are extremely fragile. Never vortex or pipet competent cells vigorously because they will lyse. Never warm competent cells to room temperature because they will start repairing their membranes and will lose their competency. Do not centrifuge the competent cells before use, because you will be unable to get them back into suspension. Set up 6 transformations (one for each ligation) as described below. 1. Set the heat block to 42˚C. 2. Label six sterile microcentrifuge tubes 1–6 (designations from ligation). Chill and keep on ice. 3. Obtain 150 µl of competent cells, keeping on ice. Keep competent cells on ice at all times. 4. Finger flick the tubes to be sure cells are in suspension, and aliquot 20 µl into each of the six chilled microfuge tubes. 5. Add 2 µl of each of the ligation mixes into the appropriate tube, being sure to add the DNA directly into the bacterial suspension. Mix the tube gently. 6. Incubate on ice 5 minutes. 7. Heat pulse the cells for 30 seconds at 42˚C. The duration and temperature of the heat pulse is critical.
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8. Incubate tubes on ice 2 minutes. 9. Add 80 µl of room temperature SOC broth or LB broth per tube and shake for 45 minutes at 37˚C. 10. Label 6 LB/kan plates with your lab day and station number and number them 1–6 (to the individual ligations). 11. Mix the cells in the tube gently, then plate 100 µl from each tube onto LB/kan plates that are labeled 1–6A. Each plate should also be labeled with your lab day and station number. Always label the backs of Petri dishes rather than the lids to avoid mixups. 12. Place the plates inverted (upside down) in the 37˚C incubator overnight.
C. Electrophoresis of Ligation Reactions This can be started during the 45 minute transformation incubation. While the transformation of the control ligations is a much more sensitive method to detect recircularization of vector (or vector that was not digested to completion), gel electrophoresis of the ligation products will give us a more rapid answer. Unlike linear DNA, circularized plasmid DNA has secondary structure, and this secondary structure affects the movement of the DNA through the gel. Therefore, circular plasmid DNA will often run at an unpredictable molecular weight. Additionally, it can run as several bands representing multiple conformations, especially if it has been digested and re-ligated. Add 1 µl of loading dye to samples 2–7 before loading into wells. 1. 2. 3. 4. 5. 6. 7.
10 µl 1 kb ladder (premixed with loading dye) 10 µl ligation 1 10 µl ligation 10 µl ligation 3 10 µl ligation 4 10 µl ligation 5 50 ng egfp insert (if you have it left over)
Run the gel as previously described. Figure 6.2 represents what your gel may look like. If the experimental ligations work, you will typically see the band corresponding to cut vector disappear, and bands lining up with uncut vector appearing, as well as some additional bands. This can be visualized quite well in lane 3, which represents the 1:3 vector:insert ligation. In this experiment, vector appears to have been cut to completion, because lane 4 (ligation 3) shows a single band, which appears to be linear DNA. In this experiment, it appears that lane 5 (ligation 4: vector only, plus ligase) contains circular DNA. This most likely indicates either that the stuffer fragment was not removed from the digested vector, and it was able to insert back into the large vector fragment and recircularize, or that two or more vectors were ligated together. This control suggests that not all colonies recovered from the ligation of vector and insert will actually contain insert. On your gel, if all lanes (except “ligation” 5) appear to contain linear vector, it is likely that there is a problem with either your ligase or your ligase buffer. To indisputably determine if this is the case, you could perform the following control: Digest pET-41a(+) DNA with Nco I only and run it on an agarose gel to ensure that it was cut to completion. Desalt the DNA, then set up a ligation using this singledigested vector and observe the sample on a gel. If the DNA did not recircularize, then there is a problem with your ligase, your ligase buffer, or your ligation technique. If the DNA does recircularize, then you could be confident that the ligase and ligase buffer are fine, and further trouble-shooting would be necessary.
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Fig. 6.2 Gel electrophoresis of ligations and ligation controls. Lane 1, 1 kb ladder. Lane 2, ligation 1. Lane 3, ligation 2. Lane 4, ligation 3. Lane 5, ligation 4. Lane 6, ligation 5. Lane 7, egfp insert.
REFERENCES Dugaiczyk A, HW Boyer, and HM Goodman. 1975. Ligation of EcoRI endonuclease-generated DNA fragments into linear and circular structures. J Mol. Biol. 96, 171–184.
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II Screening Transformants he next several experiments are aimed at determining which of the bacterial clones carry the egfp gene and can produce the GST::EGFP fusion protein. The DNA probe experiment will only indicate whether egfp is present. The restriction mapping experiment will confirm this and determine if extra sequences are present. Although unlikely, it is important to make sure that a recombinant molecule to be used for large-scale protein expression comprises exactly what is intended: one copy of the vector and one copy of the insert. The PCR experiment will determine if the gene is present in the correct orientation with respect to the promoter. The monoclonal antibody probe will test for protein expression, and the visualization of green fluorescence will reveal whether the recombinant protein is folded correctly. If this was your own research instead of a laboratory course, you would probably need to use one or a few of the screening techniques and would send the recombinant plasmid to a sequencing facility just to make sure that everything was correctly ligated. It is important to know and understand all of these techniques because if problems occur, it is helpful to use other techniques for trouble-shooting. Each of these techniques is used in other experiments; for example, antibodies are used in Western Blots for protein analysis and labeled DNA probes are used for Southern Blots.
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7 Colony Hybridizations he first steps in characterizing the results of a transformation experiment are to determine if any of the recovered clones contain plasmids with insert DNA, and then to confirm whether that DNA is inserted correctly and whether the protein can be made. You will use two types of probes to screen recombinant clones. To screen for the presence of the insert, the first probe will be the egfp gene isolated from pEGFPN1. You will use a digoxigenin-based method to label this DNA. To screen for the expression of the protein, the second probe you use will be a monoclonal antibody to the enhanced green fluorescent protein. This week’s lab requires an interim laboratory session because you will need to count your colonies and replica plate colonies at least one day before your regular laboratory period. The two experiments you begin during the regular laboratory period this week will be completed over the next two weeks.
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7A Interim Laboratory Session oal: Today you will count and record the number of transformants on each plate. You will also make replica plates of putative transformants from your experiment. These replica plates will be used to screen for positive clones (those that have the expression vector and the egfp insert).
G I. INTRODUCTION
Today you will prepare the templates to be used in the screening. Replica plating is the technique by which each colony/clone is inoculated onto multiple plates according to a numbered scheme. This method allows each clone to be tested by a variety of assays, while retaining a master plate from which the original clones can be picked. Although one plate can be used both as the master and as the template for filter hybridizations, the risk of cross-contamination between colonies is greater. We only need to screen on the order of tens of colonies, so we will pick each colony individually and inoculate replicas using a toothpick. In cases where hundreds or thousands of colonies must be replicated, a sterile velvet stamp may be touched to the original plate and stamped onto multiple blank plates to grow replicas. The individual fibers of the velvet act as tiny inoculating needles.
II. LABORATORY EXERCISES A. Counting Transformants Count and record the number of transformants on each of your ligation/transformation plates. You should have many colonies on the plate from the control ligation/transformation 5, and several colonies from your experimental ligations 1 and 2. You should have very few, if any, colonies on the plates where the bacteria were transformed with ligation 3 or 4. Why? Determine whether your controls gave you the expected results. If they did not, what technical problem could have occurred? Would you still screen the colonies found on plates where you expect to see positive results (vector ligated with insert)? If you see numerous colonies in ligation 3 and/or 4, it may be difficult to find transformants that have incorporated insert in ligations 1 and 2. Why?
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B. Replica Plating We will be doing triplicate replica plates to analyze potential positive clones from your vector-insert ligations. 1. Obtain three plates of Luria-Bertani/kanamycin (LB/kan). One will be used for the DNA probe experiment, one for the antibody probe experiment, and the third will be retained as a master. IMPORTANT: Be sure to label all three plates with your lab day and your station number. 2. Adhere grid stickers to the backs of three fresh LB/kan plates. If grid stickers are not available, place the three fresh LB/kan plates on the sheet containing the Petri plate grids (Figure 7.1) and use these as your template. 3. Obtain a plate containing pAD1 (positive control) and a plate containing pET41a(+) (negative control). Using a sterile toothpick, pick up cells from a culture of the positive control (pAD1) by lightly touching a single colony. Lightly drag the toothpick in the shape of a + in square 28. Repeat the dragging motion for all three plates (there is no need to pick up more cells between plates). Discard the toothpick into a biological waste container, and take a new toothpick. Repeat with pET-41a(+), but make the streak in the shape of a -(for negative control) in square 29 for all three replicas. You are putting the controls toward the center of the plate, because the colony hybridizations tend to be most accurate in the center and least accurate at the edges of the plate. It is critical that the controls are accurate, otherwise none of the data may be analyzed. 4. Pick up cells from your 1:1 and 1:3 transformation plates. Lightly drag the toothpick in a diagonal direction in square 1 on the first plate and then, with the same toothpick, in an identical manner on the second and third plates. To facilitate interpretation of the results, the pattern of streaks should not be symmetrical. Try not to stab the toothpick into the medium. 5. Using a different autoclaved toothpick for each colony, pick up cells from your plates of transformants. Be sure only to pick well-isolated colonies—not colonies that are touching each other. 6. Each group should pick all of their transformants from ligation 1 and 2 (up to 48 recombinant clones will fit on the plate). If you have fewer than 20 colonies, pick 6 colonies from the “mixed unknowns” plate. The mixed unknown plate contains a mixture of positive clones and clones with vector only. You may also borrow a plate from another group—you can re-pick colonies they have already picked. Be sure to record in your notes which plate each clone came from. Do not pick colonies from ligations 3–6—these will all be negative. 7. Ordinarily you would incubate the plates in a 37˚C incubator overnight. Instead, stack your plates, wrap colored tape around them (once!), and store them at 4˚C in designated drawers in the refrigerator (be sure to label your lab day and station number and that the plates are in the correct drawer or they may not be moved to the incubator!). We will come in the night before the next session and transfer them to the 37˚C incubators, so that the plates will be warm and free of condensation when we make replicas from them.
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7B Colony Hybridization: DNA Probe oal: You will screen your putative clones using a digoxigenin-labeled DNA probe to determine which ones have the egfp gene inserted. The probe corresponds to the egfp gene. We know that colonies growing on kanamycin contain plasmid DNA, but not all of the plasmids have the egfp insert. This procedure is the same as that used for screening cDNA or genomic libraries. The DNA encoding the gene of interest is amplified by PCR using a mixture of deoxynucleotides, one of which has been previously labeled with digoxigenin. Hybridizing the probe to filters lifted from plates containing bacterial colonies is used to detect positive clones that contain the sequences homologous to the probe. When screening a library made from the entire genome of a microorganism for a colony containing a single gene, a success rate of 1 positive in 10,000 screened is typical, depending on the genome size. In the present experiment, the frequency of positives should be much higher. If your cloning worked well, you may see a 50% or higher frequency of positive clones. Because this experiment involves an overnight hybridization and several washes, it will be done over a span of two weeks.
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I. INTRODUCTION Probing colonies with a labeled DNA probe confirms the presence of a specific DNA sequence. This technique is an excellent tool for screening random genomic and cDNA libraries for specific genes because DNA probes are easy and inexpensive to make, either by restriction digestion of cloned DNA or by PCR. DNA probing is also useful if you have only limited knowledge of the DNA sequence you are looking for; you can do “low-stringency” hybridization using a heterologous probe (DNA from another, related organism as the probe). While colony hybridization with a DNA probe does confirm the presence of a specific DNA sequence, it has limitations in that it does not give information about the orientation, location, or reading frame of the sequence in question. The principle of DNA hybridization relies on both the DNA being probed (the sample DNA) and the DNA probe itself being denatured (single stranded). If both the probe and the sample DNA are single stranded, then the probe will hydrogen bond with the sample DNA if the sequences match. Probes can be encouraged to bind to either perfectly matched DNA sequences, or to similar, but not perfectly matched DNA sequences. Experiments where only perfectly or nearly perfectly matched hybridizations are desired are performed under the high stringency conditions of high temperature and low salt in the hybridization and washing buffers. Experiments where less than perfect matches are desired, such as experiments using
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a heterologous DNA probe, hybridizations are performed under reduced stringency (lower temperature and higher salt). In addition to binding the sample DNA of interest, the DNA probe must itself be detectable. The classic method is to use a radiolabeled DNA probe and then to detect the probe using X-ray film. Radioactivity, however, is an environmental and safety hazard and should be avoided whenever possible. In recent years, enzymelinked assays have replaced radiolabeled probes in all but the most sensitive experiments. We will make use of a digoxigenin-labeled (dig-labeled) probe, followed by detection with a horseradish peroxidase-conjugated dig-specific antibody. This method may be used for either colony hybridizations, as we will perform, or for a Southern Blot, where DNA is purified, cut with restriction enzymes, separated on an agarose gel, denatured, and then transferred to a nylon membrane. The main principles of hybridization and detection using a digoxigeninlabeled probe are depicted in Figure 7.2. The unknown denatured DNA is immobilized on a nylon membrane by UV cross-linking. The membrane is then hybridized with the dig-labeled probe. The probe will only hybridize with DNA that contains a matching sequence. Therefore, in our experiment, the probe will anneal to plasmid DNA that received the egfp insert, but not vector-only DNA. Following the hybridization, the membrane is washed to remove unbound probe and incubated with digoxigenin-specific antibody conjugated to the horseradish peroxidase (HRP) enzyme. After a second wash, the colorimetric HRP substrate chloronaphthol is added. Chloronaphthol is cleaved by the HRP enzyme, and the product of the enzymatic reaction is a purple precipitate. Therefore, areas on the membrane where DNA containing the egfp gene is located will be purple in color, and areas that do not contain the egfp gene will show no signal. Using an egfp dig-labeled probe, we can determine which of the transformants contain plasmid DNA with the egfp insert. It should be noted that other enzyme/substrate combinations can be used for detection. A common example is the use of a digoxigenin-specific antibody conjugated to the alkaline phosphatase (AP) enzyme followed by incubation with Disodium 3-(4-methoxyspiro{1,2-dioxetane-3,2′-(5′-chloro)tricyclo[3.3.1.13,7] decan}-4-yl)phenyl phosphate (known as CSPD). Luminescence is produced when alkaline phosphatase cleaves CSPD, which is detected by exposure to X-ray film. Labeling the DNA probe with digoxigenin is fairly simple. DIG-11-dUTP is used instead of dTTP as a substrate for Taq DNA Polymerase during the polymerase chain reaction (PCR). Incorporation of digoxigenin allows the synthesis of labeled DNA probes; the digoxigenin is used in a later step as a molecular “flag” for detection. The ratio of dTTP to DIG-11-dUTP should be 2:1 for highly efficient probe labeling, suitable for single copy gene detection. The reason DIG-11-dUTP is not used to substitute for all of the dTTP is that steric hindrance of DNA synthesis due to the slightly bulkier digoxigenin could occur. Using only 1/3 DIG-11-dUTP still allows for sufficient labeling across the entire DNA molecule. While preparing the labeled DNA probe, a control PCR reaction should be run. The control reaction should contain identical reagents to the labeled probe, but without DIG-11-dUTP. When the labeled reaction is run side by side on an agarose gel with the control reaction, the labeled gel should have a slightly higher molecular weight due to the extra weight of the digoxigenin molecules (Figure 7.3). The experiment works as follows: The DNA on the nylon membrane is denatured (made single stranded), and then hybridized with excess single-stranded probe under conditions that promote annealing of the probe with complementary DNA on the membrane. Excess probe is washed away under low salt, relatively high temperature conditions (65˚C for high stringency) to remove probe that hydrogen bonded with similar, but not identical DNA sequences. Probe is then detected as described above. Figure 7.4 shows the steps of the colony hybridization with the egfp probe experiment.
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Lab Session 7B DIG molecules 1. DNA probe hybridization
DIG-labeled egfp probe
pET-41a(+) with egfp
pET-41a(+)
HRP
Horseradish peroxidase-conjugated digoxigenin-specific antibody
HRP
P
HR
2. Antibody detection
HRP
3. Detection of HRP using chloronaphthol
chloronaphthol HRP
HRP P HR
Fig. 7.2 The principle of DNA hybridization and detection. Panel 1: the singlestranded dig-labeled egfp probe binds only to the DNA of transformants that have the egfp insert. Panel 2: the anti-dig-HRP antibody conjugate binds to the dig-labeled DNA probe that is annealed to positive clones. Panel 3: chloronaphthol is cleaved by HRP, leaving a purple precipitate on the areas where the positive clones were lifted.
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Fig. 7.3 Agarose gel of the dig-labeled egfp probe versus the same sequence, not labeled. Lane 1, 1 kb ladder; lane 2, labeled probe; lane 3, unlabeled control.
Colony Hybridization with DNA Probe
replica plate
transformants on LB/kan
transfer to nylon membrane
+ pAD1, - pET-41a on LB/kan
hybridize w/ labeled DNA probe
degrade protein
membrane with DNA imprint
lyse cells and denature DNA righ on the membrane. UV crosslink DNA t membrane bacterial imprint on nylon membrane
stringency washes
develop
positives “light up” negatives do not
Fig. 7.4 Flowchart of the colony hybridization with a DNA probe experiment.
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Lab Session 7B
II. LABORATORY EXERCISES A. Colony Hybridization with an egfp DNA Probe: Part 1 Wear NON-POWDERED gloves. Oils from your fingers will inhibit transfer to the nylon membrane. Label your blot in PENCIL with your lab day and station number. 1. Notch the edge of a nylon membrane for orientation. It is a good idea to cut two distinct notches in the membrane so that there is no confusion later when trying to align the membrane with the plate. Cutting a small triangle and a small semicircle next to each other on the edge of the membrane works well. 2. Label the edge of the membrane with your lab day and station number. 3. With forceps, lay the membrane onto the plate, labeled side TOWARD the plate (that way, you will know that the labeled side is the side that lifted the colonies for future steps). Do not drag or reposition the membrane. 4. Mark the bottom of the plate to show the position of the filter paper notches with a Sharpie. 5. Wait 5 min for colonies to adsorb to the filter, and then carefully remove the filter without dragging it across the plate. 6. Wrap the plate in Parafilm, label it as the DNA probe plate, and save it in the refrigerator for reference. 7. Obtain a 1-foot-long piece of plastic wrap. Spread on a flat surface of your lab bench. 8. Obtain aliquots of denaturing, neutralizing, and 2XSSC solutions. 9. On the plastic wrap, make 2 separate pools of denaturing solution (about 0.75 ml) about 6 inches apart. 10. Incubate the membrane in one puddle, colony side up, for 5 minutes. 11. Using forceps, move the membrane to the second puddle and incubate 5 min. 12. Make 2 more puddles with 0.75 ml neutralizing solution. Incubate the membrane colony side up for 10 min in each puddle. 13. Place the membrane in a puddle of 2XSSC, and incubate 10 min. 14. UV cross-link in the Stratalinker. UV cross-linking permanently immobilizes the DNA on the membrane. If this step were bypassed, the DNA would wash off of the membrane during the prehybridiation, hybridization, and wash steps. 15. Place the membrane in a Petri dish with 0.5 ml proteinase K, incubate 1 hr at 37˚C. Proteinase K digests the cellular proteins. 16. Sandwich the membrane between 2 moistened sheets of 3MM paper and apply a slight pressure. Change to dry 3 MM paper and hand the sandwich in to instructor to freeze at −20˚C until a later date.
B. Labeling of DNA Probe by PCR Using Digoxigenin-11-dUTP Your instructor may prepare the labeled probe for you if time does not allow you to do so. Synthesis of Probes Primers corresponding to the egfp gene have been designed to amplify a 700 bp fragment by the polymerase chain reaction (PCR). Taq DNA polymerase will be used to incorporate digoxigenin into the egfp DNA to make the probe for hybridization with the nylon membranes.
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The following nucleotide concentrations are recommended for digoxigeninlabeling a DNA probe by PCR: 70µM DIG-11-dUTP, 130 µM dTTP and 200 µM dATP, dGTP, dCTP each. These probes will be incubated with the membranes where they will hybridize to clones with the egfp gene. The same probe could also be used to single copy genes in a Southern Blot of genomic DNA. To make 100 µl of DNA probe, follow the table below. You may need to make dilutions of some of the reagents. You will prepare both a dig-labeled DNA probe and a nonlabeled DNA control.
Stock Solutions dATP 100 mM dCTP 100 mM dGTP 100 mM dTTP 100 mM dUTP-dig 1mM Primer 1* 100pmol/µl Primer 2** 100pmol/µl PCR buf + Mg 10× stock Template pEGFP-N1 450 ng/µl Taq polym. Water
Desired Concentration for Probe 200 µM 200 µM 200 µM 130 µM 70 µM 1 pmol/µl 1 pmol/µl 1× 4.5 ng/µl
Amount to Add to 100 µl Probe 0.2 µl 0.2 µl 0.2 µl 0.13 µl 7 µl 1 µl 1 µl 10 µl 1 µl 1 µl 78 µl
Amount to Add to 100 µl Nonlabeled Contro 0.2 µl 0.2 µl 0.2 µl 0.2 µl 0 µl 1 µl 1 µl 10 µl 1 µl 1 µl 78 µl
*
The sequence of primer 1 (the forward primer) is CTTGTACAGCTCGTCCATGC. The sequence of primer 2 (the reverse primer) is AGAGTCCCATGGTGAGCAAG.
**
Cover the reaction in the microfuge tube with mineral oil. The mineral oil acts as a vapor barrier to prevent water from evaporating and changing the ionic strength of the PCR reaction. The total amount of mineral oil can vary as long as it forms a seal above the PCR reaction. If you are using a thermocycler that has a hot bonnet (a heated lid), there is no need to add mineral oil. Run the following program: Step 1: 95˚C, 5 min Step 2: 95˚C, 1 min Step 3: 56˚C, 1 min Step 4: 72˚C, 1 min Step 5: Repeat steps 2-4 29 times Step 6: 72˚C, 5 min to complete DNA polymerase extension of the PCR products Step 7: 4˚C, indefinitely or until you can transfer your tubes to the refrigerator or freezer. Run 5 µl of the probe and the control side-by-side on an agarose gel to ensure sufficient labeling (as described in the introduction). Be sure to include molecular weight markers.
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7C Colony Hybridization: Monoclonal Antibody Probe oal: You will screen your putative clones using an α-EGFP monoclonal antibody probe to determine which of your clones express the enhanced green fluorescent protein. This exercise will be done in 2 parts: you will prepare the blots this week, and probe them in the next lab session.
G I. INTRODUCTION
By inserting the egfp gene into the pET-41a(+) expression vector in the correct location, the orientation and reading frame of the insert should allow the EGFP::GST fusion protein to be made (expressed). Probing the colonies with the monoclonal antibody probe that binds specifically to the EGFP moiety of the fusion protein will confirm whether the recombinant fusion protein is being correctly translated. From this experiment, one can infer not only that the gene of interest is present, but also that it is in the correct orientation and reading frame. The most common method for detecting proteins on blots is to use a primary antibody specific for the protein of interest, followed by incubation with a secondary, tagged antibody that recognizes the first antibody. Our primary antibody is an α-GFP monoclonal antibody that cross-reacts with a conserved region on the closely related EGFP protein. The secondary antibody recognizes the species-specific, conserved portions of the primary antibody and is conjugated with an enzyme, horseradish peroxidase. Because the primary antibody is a monoclonal antibody (MAb) made in mouse, we will use a secondary antibody made in goat that recognizes conserved portions of mouse antibodies. We will refer to the secondary antibody as GAMP (Goat Anti-Mouse antibody conjugated to Peroxidase). The goat antibody recognizes mouse immunoglobulin G (IgG) and will therefore bind to all monoclonal antibodies made in mouse. The procedure you will use for detecting the EGFP epitope (shape of the specific portion of the protein) is similar to that for detecting the egfp DNA in that you will transfer colonies from your replica plate to a membrane (nitrocellulose instead of nylon) and then probe the membrane. There are important differences in the procedures. It is critical to induce the expression of the EGFP protein with isopropylß-D-thiogalactopyranoside (IPTG) or the protein will not be made. Protein structure is disrupted under alkaline conditions such as the denaturing solution used for preparing DNA for hybridization, so the lysis of colonies must be done under conditions that are more gentle than for DNA. Unlike nucleic acids, proteins naturally stick to nitrocellulose, so it will not be necessary to expose the filters to ultraviolet (UV) light.
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Once the colonies are lifted, the blots will be incubated in a solution with large amounts of nonspecific protein (bovine serum albumin or powdered nonfat milk) to block nonspecific antibody binding, stabilize the proteins, and saturate proteolytic enzymes that are present in E. coli. A nonionic detergent such as Triton X-100, Nonidet P-40 (NP-40), or Tween 20 is used at as low a concentration as possible to preserve epitope integrity. The nonionic detergent is included in the wash buffer to facilitate the removal of unbound antibodies. The blots are then washed without detergent to remove detergent residues that would prevent further protein–antibody interactions for the secondary antibody. Detection of the epitope will be done by incubating the blots in the colorimetric substrate chloronaphthol. Horseradish peroxidase, conjugated to the goat anti-mouse secondary antibody, cleaves chloronapthol to make a purple precipitate (Figure 7.5). A flowchart of the monoclonal antibody probe procedure is outlined in Figure 7.6.
chloronaphthol (HRP colorimetric substrate) HRP
HRP
Goat anti-mouse antibody, conjugated to horseradish peroxidase (GAMP)
EGFP-specific mAb (mouse antibodies)
proteins immobilized on nitrocellulose
chloronaphthol
GFP protein
Fig. 7.5 The principle of monoclonal antibody detection. Cellular proteins bind to the nitrocellulose membrane. The membrane is then blocked with a protein that will not interact with the primary antibody (such as casein found in skim milk). Primary antibody is added and allowed to bind to the specific epitope on the EGFP protein, and unbound antibody is washed away. Secondary antibody conjugated to horseradish peroxidase (GAMP) is added and allowed to bind to the primary antibody, and then excess (unbound) antibody is washed away. Finally, the colorimetric substrate chloronaphthol is added. Spots from transformants that expressed the EGFP fusion protein will appear purple, and clones that did not express the protein will remain colorless.
II. LABORATORY EXERCISE A. Colony Hybridization with an a-GFP Monoclonal Antibody Probe: Part 1 Because there is not enough time in a lab period to perform this entire procedure, your instructor will perform steps 1–5 for you. IMPORTANT: When working with membranes, ALWAYS wear gloves. Also, label blots in PENCIL (do not use ink).
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Lab Session 7C Colony Hybridization with mAb Probe
replica plate
transformants on LB/kan
transfer to nitrocellulose membrane
+ pAD1, - pET-41a on LB/kan
incubate w/ EGFP-specific mAb. wash unbound mAb.
wash and block.
membrane with protein imprint
induce protein expression with IPTG. lyse bacteria with chloroform. lysozyme and DNase treatment. bacterial imprint on nitrocellulose membrane
incubate w/GAMP. wash.
develop
positives “light up” negatives do not
Fig. 7.6 Flowchart of the colony hybridization with an α-GFP monoclonal antibody probe experiment
1. Prepare a filter for probing with α-GFP. Use scissors to make small notches in a nitrocellulose filter, as described for the DNA probe protocol. This will be used to orient your filter with the original plate. Label the filter with your lab day and station number. CAUTION: Never directly touch nitrocellulose or nylon membranes. Oils will inhibit the transfer of solutions to the membranes and proteins from your fingertips will bind irreversibly. Always wear gloves and manipulate the filters with blunt-ended forceps. 2. Lay the notched nitrocellulose filter in 750 µl of a solution of IPTG (20 mg/ml) in an empty Petri plate dish. The IPTG will induce synthesis of the fusion protein. 3. Place the filter on a replica plate of your transformants (labeled side toward the colonies). To prevent air bubbles, carefully drop the filter near one edge of the plate and lightly roll the membrane across the rest of the plate. Do not slide the membrane or the colonies will smear. Mark the Petri plate to show the precise location of the notches. 4. Pick up the filter and “flip” it over onto a fresh Luria-Bertani/kanamycin (LB/kan) plate so that the bacterial colonies are on top of the filter (facing up). 5. Label the plate and place the filter and LB/kan plate in a 37˚C incubator for approximately 4 hr. 6. Lyse the bacterial cells by exposing them to chloroform vapors for 30 min in the fume hood. Remove the top Petri plate and invert the filter and bottom plate over a pool of chloroform. Glass Petri plates with support mesh will be available in the chemical fume hood; pour a small amount (5–10 ml) into the glass plate, cover with mesh, and invert the bottom of the LB/kan plate with your filter directly over the chloroform. Do not remove the filter from the LB plate; allow it to remain attached to the agar during the time required for lysis.
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CAUTION: Chloroform will disintegrate plastic Petri plates on direct contact; make sure that the chloroform aliquoted for bacterial lysis is in a glass container and that a support mesh or tape is used to prevent direct contact of the inverted plate. Chloroform should be aliquoted using a glass pipet or Erlenmeyer flask. 7. Remove your filter from the hood. Separate the filter from the agar and wash the filter colony side up for 30 min in a Petri plate containing 7.5 ml of lysis buffer [blocking solution with lysozyme (40 µg/ml) and pancreatic DNase (1 µg/ml) to remove bacterial debris. 8. Incubate the MAb filter in 10 ml of blocking solution without lysozyme or DNase for 30 min (or overnight at 4˚C if desired). Use a platform shaker to gently swirl the solution. 9. Rinse in l× wash buffer with IGEPAL (a mild detergent) plus 0.1% nonfat powdered milk for 30 min at room temperature. 10. Rinse in l× wash buffer with 0.1% nonfat powdered milk for 15 min. 11. Place blot on Whatman paper and allow it to dry for 5 minutes. Sandwich the blot between 2 squares of fresh Whatman paper and turn them in to the instructor or TA. The blots will be frozen and we will continue with this protocol in another lab session.
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8 Completion of Colony Hybridization with DNA Probe oal: This week you will complete the colony hybridization with the egfp probe and analyze the results. Note to instructor: It is a good idea for the students to prepare a fresh replica plate on LB/kan medium to ensure live, healthy cells for upcoming experiments. Students should use the one of the three replica plates that was not used for the DNA or mAb colony lifts. It should still be pristine. Incubate the fresh replica plate overnight at 37˚C, and then refrigerate until use.
G I. INTRODUCTION
Review the introduction from Lab Session 7B.
II. LABORATORY EXERCISE A. Colony Hybridization with an egfp DNA Probe: Part 2 Prehybridization Your instructor will do this the day before your lab, or he/she may ask you to come in the day before your lab. 1. Place the blot in a hybridization tube with 10 ml of hybridization buffer. The labeled side should be facing the interior of the tube so that it will be continuously washed with solution. Incubate at 65˚C for 2 hours in the hybridization oven. We will be placing 2–3 blots per tube. Be sure blots do not overlap each other (and check periodically during incubation), and be sure to place the colony side on the inside, so that the side away from the colonies is against the glass. The presence of detergent will make the glass hybridization tubes slippery. Use care when handling the tubes. Hybridization Your instructor will do this the night before your lab because it needs an overnight incubation. He/she may ask you to come in to assist. Probe aliquots and hybridization tubes will be shared between 2–3 groups. 1. Obtain an aliquot of labeled probe, place in a plastic holder and incubate for 10 minutes in a 95˚C (or hotter) water bath. Screw cap tubes should be used
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because flip-top microfuges may pop open in the boiling water bath. If you are using tubes with flip-top lids, use a special lid-lock to ensure that lids remain closed. 2. Place the tube immediately on ice/ethanol to prevent reannealing of the denatured probe. 3. Spin briefly in the microfuge to bring the contents to the bottom of the tube if necessary, then place back on ice. 4. Dilute 20 µl of probe into 10 ml of hybridization buffer. This is your hybridization solution. Preheat this solution to 65˚C. 5. Decant the prehybridization solution from the blots, and add 10 ml hybridization solution containing probe to the hybridization tube. 6. Incubate overnight in the hybridization oven at 65˚C. Stringency Washes This is where you will begin when you come to lab this week. 1. Pour the hybridization buffer with probe into a 15-ml conical tube labeled with the date, your lab station, and “egfp dig-dUTP PCR-labeled probe” and freeze at −20 C. This solution is stable for 1 year and can be reused. 2. Move the blot to a fresh Petri dish and wash twice by adding 30 ml of 2 × wash solution and incubating at room temp (5 min per wash). 3. Wash twice in 30 ml 0.5 × wash solution at 65˚C (15 min per wash) Detection All incubations should be performed at room temperature with agitation. 1. After the hybridization and stringency washes, rinse the membrane briefly (1–5) min in Washing buffer. 2. Incubate for 30 min in 15 ml of Blocking solution. 3. Dilute the anti-dig-HRP antibody by adding 3 µl into 15 ml Blocking solution. This is your Antibody solution. 4. Incubate for 30 min in 15 ml Antibody solution. 5. Wash 3× 10 min in 20 ml l × wash buffer + 0.1% nonfat powdered milk. 6. After the last wash, incubate the filter in 7.5 ml of peroxide stain for 15 to 20 min at room temperature. A purple color should develop. 7. After a deep purple color develops on the positive control, wash the filter in two changes of distilled water. 8. Dry the filter on paper towels. Keep in the dark to maintain color. 9. Analyze your results by comparing your blot with the original Petri dish and aligning the positive and negative controls and the notches cut into the membrane. Record which transformant numbers are positive and which are negative as a table in your notebook.
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9 Characterization of Recombinant Clones n this laboratory session, you will complete the colony hybridization with the monoclonal antibody probe experiment, as well as begin a PCR screen and a fluorescence visualization assay.
I
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9A Completion of Colony Hybridization with mAB Probe oal: In this exercise, you will complete the colony hybridization with an EGFP-specific monoclonal antibody probe and analyze the results.
G I. INTRODUCTION
Review the introduction from Lab Session 7C.
II. LABORATORY EXERCISE A. Colony Hybridization with an a-GFP Monoclonal Antibody Probe: Part 2 1. Retrieve your blot from the freezer and rinse in l× wash buffer with 0.1% nonfat powdered milk for 15 min in a clean Petri dish. 2. Incubate in 7.5 ml of blocking solution with anti-GFP diluted 1:1000 (7.5 µl anti-GFP in 7.5 ml blocking solution). Shake at room temp for 1 hr. 3. Wash the filters for 10 min in each of the following buffers. After each wash, discard the wash solution in the sink, then invert the Petri plate and drain the remaining wash solution on paper towels. Then add 10 ml of the next solution. ● ●
●
Wash 1 (10 ml): l× wash buffer + 0.1% nonfat powdered milk Wash 2 (10 ml): l× wash buffer + 0.1% nonfat powdered milk + 0.1% IGEPAL* Wash 3 (10 ml): l× wash buffer + 0.1% nonfat powdered milk
4. Add 7.5 ml of goat anti-mouse antibody conjugated to horseradish peroxidase (GAMP) diluted 1:500 in antibody blocking solution (15 µl of GAMP plus 7.5 ml of blocking solution). Incubate the plate for 1 hr on a platform shaker at 40 rpm, room temperature. 5. Detergents inhibit peroxidase activity. Therefore IGEPAL is omitted in the following steps. Wash the plate three times (10 min each) with the following: ● ● ●
Wash 1 (10 ml): l× wash buffer + 0.1% nonfat powdered milk Wash 2 (10 ml): l× wash buffer + 0.1% nonfat powdered milk Wash 3 (10 ml): l× wash buffer + 0.1% nonfat powdered milk
*
IGEPAL is a nonionic, non-denaturing detergent that is also known by the following names: Nonidet P40; NP 40; IGEPAL CA-630; and Nonylphenyl-polyethylene glycol.
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6. After the last wash, incubate the filter in 7.5 ml of peroxide stain for 15 to 20 min at room temperature. A purple color should develop. 7. Wash the filter in two changes of distilled water. 8. Dry the filter on paper towels. Keep in the dark to maintain color. 9. Analyze your results by comparing to your original Petri dish. Record which positives correspond to which transformant number and include the information in your laboratory notebook.
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LAB SESSION
9B PCR Screening oal: This PCR screen will confirm whether clones have the egfp insert, and whether it is in the correct orientation in the vector. You will begin this experiment today and analyze the results next week.
G I. INTRODUCTION Polymerase Chain Reaction
In order to understand the PCR screening technique applied in this laboratory session, a general understanding of the polymerase chain reaction is required. The polymerase chain reaction (PCR) was developed by Kary Mullis in 1983 (Mullis, 1990). It has simplified many procedures in molecular biology and made possible countless new techniques. The PCR uses a logarithmic process to amplify DNA sequences. A thermostable DNA polymerase derived from the bacterium Thermus aquaticus (Taq), which grows in hot springs at 75˚C, is used in repeated cycles of primer annealing, DNA synthesis, and dissociation of duplex DNA to serve as new templates. The theoretical amplification of template DNA, assuming reagents are not limiting and the enzyme maintains full activity, is 2n where n is the number of cycles. After 30 cycles of PCR from a single template, 1 × 109 new DNA molecules could be synthesized. A typical PCR cycle consists of the following steps: 1. Denature DNA (94˚C) 2. Anneal primers to template (using the melting temperature of the primers ~60˚C for a typical 20-mer) 3. Synthesize DNA (72˚C) 4. Repeat steps 1–3 30 times DNA primers are complementary to the ends of the sequence to be amplified and are oriented in opposite directions. In other words, the two primers must flank a DNA region, with one primer annealing to the sense strand and one to the antisense strand, with the primers facing inwards toward each other (Figure 9.1). Both primers are necessary for exponential amplification to occur. Primers are chemically synthesized on an instrument called an oligonucleotide synthesizer. The annealing temperature of the primers can be calculated from the formula for the melting temperature (Tm) of DNA molecules shorter than 50 bp: Tm = (4)(number of GC pairs) + (2)(number of AT pairs). In designing primers, remember that DNA synthesis occurs in a 5′ to 3′ direction. The most critical nucleotide for successful amplification from a primer is, therefore, the one at the 3′ terminus.
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Part II. Screening Transformants reverse primer 5' 5'
3'
sense strand
3'
5'
antisense strand
5' forward primer
Fig. 9.1 Orientation of PCR primers in relation to target DNA. The forward primer anneals to the 3′ end of the lower strand. When the forward primer is extended, a copy of the upper strand is created. The reverse primer anneals to the 3′ end of the upper strand. When the reverse primer is extended, a copy of the lower strand is created. If the upper strand corresponds to the sense strand, the forward primer creates a copy of the sense strand, even though it binds to the 3′ end of the antisense strand. By convention, the sequence of a gene refers to the mRNA-like strand. The template used by RNA polymerase during transcription is the antisense strand of a gene. This convention makes it easier to conceptualize sequence domains and correlate them with protein motifs.
The ingredients necessary for the polymerase chain reaction to take place are: ● ● ● ● ● ● ●
Template DNA Forward primer Reverse primer Nucleotides (dATP, dCTP, dTTP, dGTP) Taq DNA polymerase Buffer Magnesium (necessary for enzyme activity)
For an excellent review of PCR, visit the website http://www.dnalc.org/ ddnalc/resources/shockwave/pcranwhole.html (Dolan). PCR has numerous applications, from cloning genes, to synthesizing labeled DNA probes, to forensic investigations. In this transformant screening technique, we will use PCR to confirm the existence of the insert in the expression vector. Because we inserted the egfp gene using the method of forced cloning, we are confident that the gene could only have been inserted in the correct orientation. This screening technique, however, is also able to confirm the correct orientation of an insert in cases where that is an issue (when the vector is cut with only one enzyme). The key to this experiment is the use of specially designed PCR primers. As in any PCR reaction, the two primers flank a DNA region, with one primer annealing to the upper strand and one to the lower strand, with the primers facing inwards toward each other. The special feature of these primers is that one primer must anneal to insert sequence and one must anneal to vector sequence. If the insert is present and in the correct orientation, a PCR product that is the size of the flanked DNA sequence will be produced (in our case, this is 1500 bp). The product can be visualized by running a small amount of the PCR reaction on an agarose gel. If the insert is not present in the vector, the primer that was designed to anneal to the insert will be unable to bind, and no PCR product will be made. If the insert was cloned into the vector in the incorrect orientation, both primers will anneal to the same strand of DNA in the same orientation, and exponential amplification will be unable
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to occur, so no PCR product will appear on an agarose gel. We can therefore ascertain that if a PCR product is visible on an agarose gel, then the insert is present and in the correct orientation (Figure 9.2). Remember that DNA synthesis can only occur in a 5′ to 3′ direction. What would you hypothesize the result would be if an incorrect insert was cloned into the vector?
primer 1
primer 2
5'
3'
3'
5' correct orientation = product
5'
3'
3'
5' no insert = no product
5'
3'
3'
5' incorrect orientation = no product (shouldn’t occur with forced cloning)
Fig. 9.2 Screening transformants by PCR. In the first possibility, egfp is inserted in the correct orientation in the vector. The primers anneal in the correct position and 1500 base pair PCR product is made. In the second possibility, egfp is not present, and so Primer 1 cannot anneal and no PCR product is made. In the final scenario, egfp is present in the reverse orientation. Both primers bind, but in the same orientation, to the same strand. No PCR product is made.
II. LABORATORY EXERCISE A. Polymerase Chain Reaction Screen for Recombinant Clones: Part 1 You will complete this protocol during your 1 hr incubation with monoclonal antibody from the antibody probe protocol. The PCR procedure you will follow is similar to that for amplifying isolated DNA except that the first step, 95˚C degrees, takes longer in the first cycle because we have to not only denature the DNA, but also ensure that the bacteria are lysed by the heat to release the DNA. At 95˚C, most cellular enzymes that degrade DNA become heat inactivated allowing PCR to occur without a clean DNA preparation. Each station will receive one tube of master mix, or you may be asked to make your own master mix following the chart below. 1. The recipe for the master mix is shown in the following table. Students will do 8 reactions, 20 µl per tube.
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per 180 µl
Item Water A T C G Primer1 (pad1sense)* Primer2 (pad1anti)* 10× Buffer with Mg Taq polymerase
155.2 µl 0.36 µl 0.36 µl 0.36 µl 0.36 µl 0.9 µl 0.9 µl 18 µl 3.6 µl
*The pad1sense primer anneals to the 3′ portion of the sense strand of the egfp gene. The sequence of the pad1sense primer is CTTGTACAGCTCGTCCATGC. The pad1anti primer anneals to vector sequence (specifically, sequence in the middle of the antisense strand of the gst gene). The sequence of the pad1anti primer is CAAGCTACCTGAAATGCTGA. The predicted PCR product is 1500 base pairs in size.
2. Aliquot 20 µl of reaction mix into eight small strip tubes. You will screen five putative transformants, and include a no-template DNA control to test for contaminant DNA, a pAD1 positive control, and a pET-41a(+) negative control (8 total). 3. Use a sterile yellow pipette tip (wooden toothpicks contain an inhibitor of PCR) to pick up a small amount of bacterial growth from a putative transformant from your freshest master plate. Swirl the tip in PCR reaction mix. Label the PCR tube with the number on your master plate. Repeat with a different clone for each tube (including controls). For this experiment, LESS IS MORE. Pick up a very small amount of bacterial growth, about the amount that would fit on the head of a pin. Putting too much bacteria in the PCR reaction will result in degradation of PCR product. 4. Run the following program: Step Step Step Step Step Step ucts
1: 2: 3: 4: 5: 6:
95˚C, 10 min (denature) 95˚C, 1 min 56˚C, 1 min 72˚C, 1.5 min Repeat steps 2–4, 29 times 72˚C, 5 min to extend all unfinished prod-
You will analyze the results next week, using gel electrophoresis.
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9C Visualization of Green Fluorescent Protein: Part 1 oal: You will replica plate your transformants (and positive and negative controls) onto LB medium that contains kanamycin and IPTG. This will allow for visualization of green fluorescence in positive clones. You will also prepare a replica plate onto LB/kan medium so that you will have a fresh master plate.
G I. INTRODUCTION
As discussed in Lab Session 7c, IPTG acts to de-repress the T7/lac promoter on expression vector pET-41a(+). By incorporating IPTG into the growth medium, clones that can express the enhanced green fluorescent protein will appear slightly green even under ambient light. Positive clones will fluoresce bright green when UV light is shone on them.
II. LABORATORY EXERCISE A. Green Fluorescence Assay and Preparation of a Fresh Master Plate You will use your freshest master plate to replica plate onto the LB/kan/IPTG plate, as well as onto a new LB/kan plate. 1. Retrieve your master plate from the refrigerator. 2. Obtain 1 LB/kan/IPTG plate and 1 LB/kan plate. Adhere grid stickers to the backs and label with your lab day, station number, and today’s date. Always label the backs of Petri dishes rather than the lids to avoid mix-ups. 3. Using a toothpick, replica plate from your master plate onto the two fresh plates. Do this as you did in Lab Session 7A, using a sterile toothpick to pick a small amount of bacterial growth from each grid square on the master plate and inoculating the same squares on the two fresh plates. Be sure to change toothpicks between each clone you pick, but inoculate both fresh plates with the same toothpick (only pick up the transformant from the master plate once). 4. Tape plates shut and place in an inverted position in the 37˚C incubator overnight. Your instructor will remove these plates to the refrigerator tomorrow, and you will observe the IPTG plate next week in lab. The LB/kan plate will be used as a new master plate.
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REFERENCES Dolan DNA Learning Center: Biology Animation Library, Cold Spring Harbor Laboratory. Aug 11, 2005. Mullis, KB. 1990. The unusual origin of the polymerase chain reaction. Sci. Am. 262(4) 56–65.
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10 Further Characterization of Recombinant Clones his week you will analyze the results of the PCR screen and the fluorescence visualization assay that were started last week. You will also isolate plasmid DNA from transformants and perform restriction digestion analyses.
T
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10A Interim Laboratory Session oal: Inoculate cultures to be used to isolate plasmid DNA for restriction analysis during lab next session.
G I. LABORATORY EXERCISE
A. Inoculate Cultures for Minipreps Heavily inoculate 2 ml LB/kan cultures with four putative positive clones, one putative negative clone, and the positive control, pAD1 (6 tubes total). If you desire, choose the same transformants you chose for the PCR screen. Inoculate tubes by picking bacterial growth off the Petri dish using a sterile toothpick and then dropping and leaving the toothpick in the tube containing growth medium, being careful only to touch the end of the toothpick that is not in contact with the medium. This method does not utilize the principle of aseptic technique. It does typically work well for inoculating cultures for minipreps because the antibiotics in the medium keep most contaminants from growing, and because the inoculum is so large that it would likely out-compete any contaminant. In cases where a pure culture is critical, such as creating a freezer-stock, do not utilize this method.
IMPORTANT: Your instructor will incubate these for you the day before your lab so cultures will be fresh. Label your lab day and station number on the tubes, and place them in the designated test tube rack. Also include the transformant number on the label for each tube. Discard all old plates except for the most recent replica plate and the IPTG plate. Use the biohazard bags for disposal.
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10B Analysis of PCR Screen Results oal: You will analyze the products of the PCR screening technique performed last week by agarose gel electrophoresis. Recall that reactions that produce a 1500 bp fragment are expected only from clones that contain the egfp insert.
G I. INTRODUCTION
Review Lab Session 9B.
II. LABORATORY EXERCISE A. Gel Electrophoresis and Analysis of PCR Samples from Last Week 1. Add 2 µl of loading dye to each PCR reaction. 2. Pour a 1% agarose gel and run 15µl of PCR reactions per lane (with loading dye). In order to be able to load the molecular weight marker in an 8-well gel, leave out the no-template PCR control from gel, but do not discard until after viewing results—you will need to run it if your negative control (pET-41a(+) ) appears positive. 3. View the gel under UV light, and photograph. Positive clones will have a PCR product of approximately 1.5 kb, and negative clones will have no PCR product. Record your results in your lab notebook.
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10C Isolation and Characterization of Miniprep DNA from Potential Transformants (Restriction Enzyme Analysis of Putative Transformants) oal: You will isolate crude plasmid DNA from several transformants, then digest them with the restriction endonucleases Nco I and Not I to confirm the presence of the 0.7 Kb egfp gene. You will also perform a digest with Nco I only. This single digest will linearize the plasmid, giving rise to a single 6901 base pair fragment. A band of a larger size, or the appearance of (an) additional band(s) could indicate the presence of multiple inserts.
G I. INTRODUCTION
Whether you use blue–white screening (α complementation) or colony hybridization to identify transformants that may have an insert, you will want to isolate DNA for the final analysis. Protocols for the isolation of small amounts of DNA (minipreps) are generally quick and easy, but the resulting DNA still contains impurities. We will use a scaled-down version of the alkaline lysis procedure presented in Lab Session 2 to isolate DNA. There are several other methods that are quicker (i.e., “easy preps,” an ecologically friendly protocol that uses only one tube per sample (Berghammer and Auer, 1993)) but the alkaline lysis method is the most reliable. For analysis by digestion with restriction enzymes, it is not necessary to purify the DNA on an anion exchange column. However, it should be further purified if it is to be used for cloning or sequencing. Whenever you electrophorese restricted miniprep DNA, one lane should contain uncut DNA as a reference. You may want to decrease the amount of miniprep DNA used in a restriction enzyme digest to dilute possible inhibitors. Very often, miniprep DNA will not cut to completion, but will give you the information you need about whether your insert DNA is present. However, beware of extra bands! Remember, anything can happen in a ligation; small amounts of stray DNA, as well as the insert you want, can theoretically be cloned. The bottom line in analyzing transformants is, if it matches the pattern you expect, keep it. If it doesn’t, throw it out. When analyzing restriction digestion patterns, keep the following principles in mind: ●
The DNA migrates at a rate inversely proportional to the size of the DNA fragment. From previous experience, you know that including a lane of standard molecular weight markers is critical for analysis of your DNA. Because the molecular weight 81
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●
●
●
of each of the bands in the DNA standard is known, you can use these molecular weight markers to estimate the size of unknown sample fragments. Circular supercoiled DNA travels at an unpredictable rate through the gel due to secondary structure. Therefore, the size of uncut DNA cannot be estimated using a typical DNA ladder. Multiple bands may also occur due to variations in secondary structure. The sum of the molecular weight of the fragments generated by digestion of a circular molecule equals the molecular weight of the uncut molecule. (To visualize this, think of cutting a rubber band into pieces; the total length of the pieces is the length of the rubber band.) The intensity of band fluorescence is directly proportional to the mass of the fragment. Deviations from this principle are indicative of partial digests or a doublet (two bands of the same size).
II. LABORATORY EXERCISES A. Alkaline Lysis and Ethanol Precipitation of Miniprep DNA 1. Transfer 1.5 ml of the overnight culture to a labeled 1.5 ml microcentrifuge tube. The remainder can be discarded. Repeat for each of the cultures (you should have six total, including pAD1). 2. Spin the tubes at 12,000 rpm in a microcentrifuge for no more than 30 sec (longer spins will make the pellets difficult to resuspend). 3. Remove the supernatant. Use a yellow pipette tip to remove as much supernatant as possible. The proteins in LB have a detrimental effect on restriction enzyme activity. 4. Add 100 µl of solution I and vortex until the pellet is completely resuspended. 5. Add 200 µl of freshly prepared solution II. Invert five times to mix and place on ice. 6. Leave the tubes on ice for 5–10 min (until the solution looks clear, but no longer than 10 min). 7. Add 150 µl of solution III. Invert to mix. 8. Leave the tubes on ice for 10 min. 9. Spin the tubes for 10 min in a microcentrifuge. 10. Using a 200 µl pipette tip, transfer the supernatants (the supernatant contains the plasmid DNA—do not discard!) to new, labeled tubes. Avoid transferring any white particulate matter. If necessary, to remove particulate matter, centrifuge the supernatant-containing tubes again (10 min) and transfer to a new tube. It is better to lose a little DNA yield than to carry over the particulate matter. 11. Add 2 vol (about 900 ul) of 95% ethanol (stored at −20˚C) to the supernatant. Vortex and place in freezer for 5 minutes. At this point the tubes could be stored in the freezer for extended periods of time. 12. Centrifuge the tubes for 5 min at maximum speed. 13. Remove the supernatant and discard. 14. Rinse the pellets with 1 ml of 70% ethanol by vortexing briefly and then spinning for 2 min at maximum speed. Remember—always keep the hinges of the microcentrifuge tubes pointed out. 15. Remove the supernatant and place the tube open in a Speed-Vac (Savant, Hicksville, NY) for approximately 5 min to dry. Tubes may be air-dried if a SpeedVac is unavailable. 16. Resuspend the pellet in 50 µl of TE.
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Unlike DNA purified by anion exchange, this miniprep DNA is not clean. Do not store for long periods of time at 4˚C or it will be degraded (it must be stored at −20˚C).
B. Restriction Enzyme Analysis of Miniprep DNA You will digest DNA from pET-41a(+)/egfp transformants to confirm the presence of the insert. Since we used the NcoI and Not I restriction sites to clone the egfp gene into pET-41a(+), we will use these enzymes to release the insert. We will also set up a single digest using Nco I to ensure that multiple inserts were not ligated into the vector. Follow the procedure described below to set up six double digests with Nco I and Not I and six single digests with Nco I only. It will be useful to draw a mock gel predicting the number and size of DNA fragments you would predict from the restriction enzyme digestion. To Make Master Mixes 1. You will need to set up two master mixes: one for the double digests and one for the single digests. 2. Determine the reagent volumes (enzyme, restriction buffers, water) for one enzyme digest (see Table 10.1). 3. Multiply the volumes by the number of tubes plus one, to correct for pipetting errors. 4. Combine first water, then restriction buffer, then enzyme(s) in one tube (master mix). Add RNase to the master mix to a final concentration of 20 µg/ml to degrade any RNA. Vortex your tubes and keep on ice. 5. Aliquot 10 µl of master mix to each tube. Make sure the tubes are labeled and then add 10 µl of miniprep DNA to each appropriately labeled tube. Ten microliters of miniprep DNA will have between 200 and 600 ng of DNA. Since most enzymes are sold in concentrations of about 10 units/µl, 1 µl of enzyme is more than enough for one miniprep DNA digestion. For this exercise, you will use 5 units per digestion or 0.5 µl of enzyme. This is a fivefold excess of the amount of enzyme needed to digest 1 µg of DNA in one hour. 6. Incubate the reactions for 1 hr at 37˚C. (You should have 12 tubes: 6 double digests and 6 single digests). 7. Add 1/10 vol (2 µl) loading buffer directly to the tubes. 8. Pour a 1% agarose gel. TABLE 10.1 Master Mix Calculations Master Mix for Double Digests Component Sterile water Restriction enzyme buffer 3 BSA RNase Nco I Not I DNA Total volume: *
1 tube
7 tubes
6.6 µl 2 µl 0.2 µl 0.2 µl 0.5 µl 0.5 µl 10 µl* 20 µl
To be added to numbered tubes separately (not in master mix).
Master Mix for Single Digests 1 tube 7.1 µl 2 µl 0.2 µl 0.2 µl 0.5 µl —— 10 µl* 20 µl
7 tubes
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9. Load 15 µl per well. Load 1 kb ladder, uncut pAD1*, and each of your digested miniprep DNA samples. Depending on your gel apparatus, you may not have enough wells to run all of your samples. Many gel apparati allow you to cast two rows of wells in a single gel by using two combs. If you choose to do this, make sure to add 1 kb ladder in each gel level. Run the gel until the dye-front reaches halfway. If you cast two rows of combs, do not allow the dye front of the sample in the top row to run further than the second row of wells. For the double digest, negative clones corresponding to the vector without an insert should give a single band the size of the pET-41a(+) vector, approximately 6 kb. Positive clones, which are vector plus egfp insert, should produce two bands: approximately 6 kb and 700 bp. For the single digest, negative clones should give one band of approximately 6 kb, and positive clones will give a band of approximately 6.7 kb. Clones with multiple inserts will give either a single band larger than 6.7 kb, or multiple bands, depending on the construct. Very often miniprep DNA will not cut to completion. A comparison of digested DNA with uncut DNA will tell you which bands are really extra and which ones represent refractory DNA that will not cut. If you find extra bands that cannot be accounted for, discard the transformant. Keep screening until you find a transformant that gives you the correct pattern of restriction fragments, keeping in mind that uncut DNA is not an extra band.
For the uncut pAD1, prepare the sample as follows: add 1 µl uncut miniprep pAD1 DNA to 8 µl water and 1 µl loading buffer. *
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10D Visualization of Green Fluorescent Protein: Part 2 oal: You will observe the transformants that you plated on IPTG medium using ultra violet (UV) light. Positive colonies that contain the egfp gene will glow bright green!
G I. INTRODUCTION
As discussed in Part 1, the enhanced green fluorescent protein has an excitation peak at 395 nm (UV or blue light) and emits light at 510 nm. Therefore, when you shine UV light on your positive transformants, they will appear bright green. This assay, since it is so easy to perform, will be used to judge the success of the other screening techniques you used in Laboratory Sessions 7 through 10. Most proteins are not autofluorescent. In a research lab, it is rare to be so lucky as to be trying to clone a gene that has such a simple assay for expression. For our purposes, it serves as an unambiguous positive control that can be used to determine which of the other techniques were the most reliable.
II. LABORATORY EXERCISE A. Visualization of Clones Expressing the Enhanced Green Fluorescent Protein on IPTG Plates 1. Obtain the IPTG replica plate you inoculated earlier this week. 2. Remove the lid and invert the plate (open side down) on a UV transilluminator. While viewing your transformants on the ultraviolet light box, you must wear a UV protective face-shield. 3. Turn on the ultraviolet light and view the plate. The positive control and positive clones will fluoresce bright green due to the expression of the enhanced green fluorescent protein, as shown in Figure 10.1. Record which clones were positive and which were negative in your laboratory notebook.
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Fig. 10.1 Fluorescence of E. coli expressing the enhanced green fluorescent protein. The positive control, pAD1 is inoculated in grid number 28. Clones numbered 5, 18, 23, 43, and 46 are also positive. The rest of the clones are negative.
Now that you have performed several different techniques for screening transformants, it is interesting to compare the reliability of each method. Use the following chart to do so. Leave blank the spaces for transformants that were not tested in specific assays. Keep in mind that the fluorescence test was the most reliable assay method, so use it as your reference for determining which assays gave the most consistent results.
REFERENCES Berghammer, H, and B Auer. 1993. “Easypreps”: Fast and easy plasmid minipreparation for analysis of recombinant clones in E. coli. Biotechniques 14, 524, 528.
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Transformant Number
DNA Probe
mAB Probe
PCR Screen
Restriction Digestion
Fluorescence Assay
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PA R T
III Expression, Detection, and Purification of Recombinant Proteins from Bacteria
n this portion of the course, you will learn how to purify a genetically engineered protein from bacteria. This is not an idle exercise. As practicing and would-be cell biologists, we are interested in the events that govern the life of a cell. How is the cell cycle controlled? What happens when a developing cell begins to differentiate? What are the factors that control cell growth? Why and how do cells die? If the organism is a pathogen, how is it able to infect the host? Before we can answer these questions in detail, we must know more about the actual molecules that participate in these processes. The classic biochemical solution to this problem is to take the cell apart and isolate each of its components for study. For proteins that are highly expressed in the cell, this approach is generally feasible. However, nothing short of herculean efforts have been necessary to purify protein or peptides that are present in vanishingly small quantities. This represents an important limitation since highly active gene products such as enzymes or hormones are typically present at very low levels in biological materials. With the advent of recombinant DNA techniques and the abundance of sequence information, it is often much easier to clone the gene and express it in bacteria than it is to purify the protein from the native organism. You have now finished cloning the egfp gene into the pET-41a(+) expression vector. As you will see, the enhanced green fluorescent protein (EGFP)::glutathione-S-transferase fusion protein may constitute as much as 10% of the total bacterial protein. The abundance of the fusion protein reduces the amount of contaminating protein but it is also necessary to have a method to fractionate it. The pET vectors have been engineered to allow a one-step fractionation method that can be used to make pure protein in relatively large quantities. In this part of the course, you will induce the expression of fusion protein from your plasmid clone. Induced bacterial cultures will be harvested and solubilized by treatment with the detergent sodium dodecyl sulfate (SDS) and sonication, followed by purification of the EGFP::GST fusion peptide by affinity chromatography. You will conduct protein assays to monitor the quantity of the fusion protein in different stages of the purification procedure and will visualize fluorescence to ascertain that the fusion protein is correctly folded and retains functional integrity. The purity of the final preparation will be analyzed by SDS gel electrophoresis. You will also learn an immunoblotting technique in which a monoclonal antibody will be used to detect specifically the EGFP::GST fusion protein bands on the gel.
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11 Expression of Fusion Protein from Positive Clones and Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE) and Immunological Analysis (Western Blot): Part 1 oal: This week, you (or an instructor) will induce 2-ml cultures of your clones with isopropyl-ß-D-thiogalactopyranoside (IPTG) and prepare protein samples for gel electrophoresis. IPTG must be added to the medium in order for protein expression to occur. To prevent proteolysis of the fusion protein, which can occur during stationary phase, actively growing bacterial cultures are used. If the protein is toxic, IPTG can be added a few hours before the bacteria are harvested, as previously described. Today, you will make minilysates of the cultures, run them on SDS-polyacrylamide gels, and transfer them to a nitrocellulose membrane to perform an immunoblot (also called a Western Blot). You will stain the membrane using a water-soluble dye, and then save it for next week to finish the Western Blot.
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11A Interim Laboratory Session
I. LABORATORY EXERCISE A. Inoculate Cultures for SDS-PAGE 1. Heavily inoculate three 1 ml LB/kan cultures with positive clones from the master plate that you incubated most recently. Use a separate toothpick to inoculate each of the LB/kan tubes. 2. Label the three LB/kan cultures with your lab day, station number, and the transformant number. 3. Label three empty snap-cap tubes with your lab day, station number, and transformant number to correspond with the cultures you started (you should end up with two sets of labeled tubes-one inoculated, and one empty) 4. Place the inoculated cultures and empty labeled tubes in designated racks (make certain you put it in the correct rack). The LB/kan tubes containing your inoculated cultures will be refrigerated to retard growth. The night before your lab period, they will be placed in a shaker at 37˚C. This is done to provide you with actively growing stocks for optimal protein production. The empty, labeled tubes will be used for protein induction.
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11B Expression of Fusion Protein from Positive Clones and Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE) and Immunological Analysis (Western Blot): Part 1
I. INTRODUCTION A Western Blot is similar to a colony hybridization using a monoclonal antibody probe, except that isolated proteins are first separated by molecular weight and then transferred to nitrocellulose. Sodium Dodecyl Sulphate Polyacrylamide Gel Electrophresis, or SDS-PAGE, is used to separate proteins. In SDS-PAGE, the mobility of protein molecules is determined solely by protein molecular weight, as opposed to other types of protein gel electrophoresis in which migration is dependent on numerous factors. SDS-PAGE of proteins has numerous applications, including molecular weight determination, determining sample purity, quantifying expression, immunoblotting (Western Blot), and isolating proteins for peptide sequencing or for generating antibodies. You are already familiar with DNA agarose gel electrophoresis, and SDSPAGE does share some similarities with this method. Both methods separate molecules by size, use electrical charge differences to cause migration, and both require a matrix to separate molecules by size. There are several differences between the two types of electrophoresis. 1. DNA is routinely run on agarose gels, while proteins are generally run on polyacrylamide gels. This is because proteins are generally smaller than DNA, and polyacrylamide matrices have a smaller pore (sieve) size than agarose. An exception to this is in DNA sequencing: very short DNA molecules are run on polyacrylamide gels. 2. By convention, DNA is run at constant voltage and protein is run at constant current. 3. All DNA is negatively charged, but proteins have varying charges depending on the amino acid content of the specific polypeptide and the pH of the buffer. Some proteins are positively charged, while some carry a net negative charge. 4. DNA, especially linear DNA, has little secondary structure, while proteins can be globular or linear, and can form dimers and other multimers.
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Because of #3 and #4 above, if proteins were run on a nondenaturing (no SDS) polyacrylamide gel, their migration would depend on at least three factors: size, charge, and shape. SDS PAGE allows proteins to migrate by size alone, through the use of SDS and a reducing agent. SDS is an ionic detergent that denatures (unfolds) proteins by wrapping around the polypeptide backbone forming a micelle, and thus confers a net negative charge in proportion to polypeptide length. SDS also disrupts most noncovalent bonds, thereby decreasing protein folding. A reducing agent such as β-mercaptoethanol or dithiothreitol is added to reduce cystine bonds (disulfide bonds), and further unfold the proteins. After boiling a protein sample in SDS and β-mercaptoethanol, all proteins act as negatively charged linear molecules and can be electrophoretically separated by size alone. After running the gel, it can either be stained to visualize the protein bands using Coomassie Blue, GelCode Blue, or silver stain, or the proteins can be transferred to a nitrocellulose membrane for immunoblotting (Western Blotting). In Lab Session 14, we will run an SDS-PAGE gel and stain it using GelCode Blue to visualize protein bands. In today’s lab session, we will perform a Western Blot (to be completed in the following laboratory session). The first step of this process is to transfer the proteins from the polyacrylamide gel to the nitrocellulose membrane. Total protein on the nitrocellulose membrane may be visualized at this point using the water-soluble Ponceau stain. After the proteins are transferred, probing the blot is very similar to the protocol performed in Lab Session 7c, “Colony Hybridization with a Monoclonal Antibody Probe,” except the protein on the membrane was derived from the polyacrylamide gel, rather than the colony lift. This portion of the Western Blot will be completed in the next laboratory session. The molecular weight of the GST::EGFP fusion protein can be estimated assuming the average weight per amino acid is equal to 114 Da. The gst gene is 660 bp, encoding 220 amino acids: 220 × 114 = 25,080 Da. The egfp gene is 720 bp, encoding 240 amino acids: 240 × 114 = 27,360 Da. The weight of the fusion protein can therefore be approximated as: 25,080 Da + 27,360 Da = 52,440 Da or ~52 kD You will be able to visualize this band in your positive clones using the Ponceau stain this week, and then confirm that it is indeed the correct band at the completion of the immunoblot exercise ne1xt week. In the negative clones, you may see a band of approximately 25 kD, corresponding to the GST protein alone (with nothing fused to it).
II. LABORATORY EXERCISE A. SDS-Polyacrylamide Gel Electrophoresis and Western Blot: Part 1 Gel Apparatus Preparation Note: the instructions for assembling the gel apparatus and transblot assembly are specific for the Bio Rad Mini-Protean 3 Electrophoresis unit and the Bio Rad Mini Trans-Blot Cell. If you are using equipment made by another manufacturer, your instructor will provide you with assembly and usage instructions. We are using pre-cast polyacrylamide gels, so you don’t have to pour them. If you must pour your own polyacrylamide gels in the future, keep in mind that nonpolymerized acrylamide in both the powder and liquid forms are potent neurotoxins. Great care must be taken to avoid inhalation, ingestion, and skin exposure to this substance. Solidified polyacrylamide does not pose the same safety hazard.
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1. Two gels must be run per unit, so you need to pair with another group. 2. Assemble the BioRad apparatus as demonstrated by your TA. 3. Add Tris-glycine running buffer to the top of the center buffer chamber (be sure buffer covers wells) and also to the outer chamber (only about 2 inches deep). Check for leaks. 4. Wash wells with Tris-glycine running buffer. Sample Preparation Steps 1–3 will be done for you. 1. Inoculate 1 ml of each overnight culture into 2 ml of 2× YT/kan medium containing IPTG. 2. Also inoculate positive (pAD1) and negative (pET-41a(+)) controls into 2× YT/kan/IPTG medium for each lab group. 3. Incubate at 37˚C, 3–4 hours. 4. Set heat block to 95˚C or higher (be sure heat block has water in the holes for heat transfer). 5. Harvest cultures. Spin 1.5 ml of each culture for 2 min in a microcentrifuge (use locking or screw-top microfuge tubes, if available). Also, prepare two additional samples: pAD1 and pET-41a(+) which have been grown in 2 × YT/kan/IPTG medium for you. Remove and discard supernatants, saving pellets in tube. Invert tubes over paper towels to remove residual liquids. 6. To each tube, add 50 µl dH2O and vortex until pellet is dispersed. 7. To each tube, add 50 µl of 2× sample buffer. DO NOT VORTEX TO MIX. 2× sample buffer has SDS which will foam when vortexed. Mix by gently pipetting up and down. 8. Cap the tubes firmly and use tube tabs to lock caps shut. Incubate at a minimum temperature of 95˚C for 10 minutes (be sure water is in heat-block holes— incubating in boiling water is fine). 9. Microcentrifuge the tubes for 15 minutes. 10. Carefully transfer about 40 µl of supernatant from each tube to a new labeled tube. Be careful not to transfer any pellet material. Although the samples can be stored at −20˚C, we will run them today on SDS polyacrylamide gels. Store them on ice until the gel is ready. Preparing Molecular Weight Standards We will be using the New England Biolabs (Ipswich, MA) broad range molecular weight marker (Figure 11.1). The molecular weight marker should already be mixed with loading buffer. When you are ready to load the gel, make sure the tube of molecular weight standard is securely capped and heat it for about 1 min at 90˚C. Place on ice until you are ready to load it. Loading Samples on the Gel 1. To practice, use a micropipette tip to add 2.5 µl of 2× sample buffer (containing bromophenol blue) to a lane. Do not try to expel all of the liquid in the tip: an air bubble will result, which could distort the migration of your sample, and also could force your sample out of the well. Practice until you feel comfortable, by loading 2× sample buffer. 2. Add 10 µl of MW-STD, 15 µl of the positive control (pAD1), 15 µl of the negative control (pET-41a(+)), and 15 µl of each transformant fusion protein sample (from tubes 1–3) to individual lanes according to the following scheme. Do not load sample into the first lane (you can practice in it). 2 MW-STD
3
4
5
6
7
pAD1
pET-41a(+)
Sample 1
Sample 2
Sample 3
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- 27
- 20
- 14 10-20% SDS-PAGE
Fig. 11.1
-7 - 2/3
The New England Biolabs broad range molecular weight marker.
Electrophoresis 1. Two gels will be run per apparatus: Coordinate with another group before you start. 2. Attach the electrodes to the power supply. Caution: Be sure to match the anode (red) with the positive lead and the cathode (black) with the negative lead. 3. Turn the power control knobs fully counterclockwise, to the zero setting. 4. Activate your power supply. Run your gel at 20 mA constant current. If you are running two gels on a single unit (most of you will), the total current should be 40 mA. If there are two units per power supply, increase the current according to the number of additional gels (for example, 60 mA for three gels and 80 mA for four gels). Because this is a discontinuous gel system (the running buffer and the gel buffer have a different pH and ionic strength), the current will vary during the run. Designate one person to monitor the current, adjusting the power source to maintain 20 mA (check it every 15 minutes or so.) 5. Run the gel until the bromophenol blue tracking dye reaches the bottom of the gel. Stopping Electrophoresis 1. Turn off the power supply. 2. Detach the leads from the power supply. Remove your gel from the apparatus. If the other gels connected to the power supply have not finished, turn the power supply back on, adjusting the current to the proper amperage for the number of remaining gels. 3. Pour the buffer into the sink. 4. Disassemble the gel from plates as demonstrated. 5. Cut the wells off gel after disassembling. It is possible to stain the gel to visualize the proteins at this point, but for a Western blot, the gel cannot be irreversibly stained. Instead, we will first transfer the protein to a nitrocellulose membrane, and then stain the membrane with a water-soluble dye prior to completing the Western Blot.
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Transferring to Nitrocellulose We will use the Bio-Rad mini trans-blot cell for transfer of protein from the polyacrylamide gel to the nitrocellulose membrane (Figure 11.2).
Lid
Fiber pad Filter paper Membrane Gel Filter paper Fiber pad Gel holder cassette
Electrode module
Bio-Ice cooling unit (keep frozen at −20⬚C)
Buffer tank
Fig. 11.2 Mini trans-blot cell description and assembly of parts. Copyright 2004 Bio-Rad Corporation. Permission to use these materials has been granted by Bio-Rad Laboratories, Inc. This permission is solely for inclusion in Manipulation and Expression of Recombinant DNA: A Laboratory Manual.
1. Prepare 1 liter 1× transfer buffer by mixing 200 ml 5× transfer buffer with 200 ml methanol and 600 ml distilled water. 2. Obtain a glass dish, 2 sheets of filter paper and 1 sheet nitrocellulose. Be sure to handle everything only when wearing gloves. 3. Label nitrocellulose on top left in pencil. 3. Pour a small amount (about 1 cm deep) of 1× transfer buffer into dish. 4. Soak nitrocellulose membrane, gel, fiber pads, and filter paper 10 minutes. Fiber pads are not disposable. Be sure not to allow the nitrocellulose to touch the gel at this point. 5. Prepare a gel sandwich as described below and depicted in Figure 11.3. Under no circumstances should you shift the position of the nitrocellulose on the gel. a. b. c. d.
Place the cassette, gray side down, on a clean surface. Place one prewetted fiber pad on the gray side of cassette. Place a sheet of filter paper on pad. Place the gel on the filter paper (remove bubbles by gently rolling a glass tube over it).
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e. Place the prewetted nitrocellulose membrane on the gel (labeled side down at top of gel; remove bubbles by gently rolling a glass tube over it). f. Place a filter paper on the membrane (remove bubbles by gently rolling a glass tube over it). g. Add the last fiber pad and close the sandwich.
Fiber pad Filter paper Membrane Gel Filter paper Fiber pad
Fig. 11.3 Close-up of “gel sandwich.” Copyright 2004 Bio-Rad Corporation. Permission to use these materials has been granted by Bio-Rad Laboratories, Inc. This permission is solely for inclusion in Manipulation and Expression of Recombinant DNA: A Laboratory Manual.
6. Place the cassette in the module. The gray side of the cassette should face the black side of the module, and the clear side of the cassette should face toward the red side of the module. 7. Add an ice pack unit. 8. Add transfer buffer to the top (if you run out, use the transfer buffer you used for soaking your gel, etc.). 9. Put on the lid of the assembly so that red matches red and black, black. 10. Turn on the voltage for 1 hour, 100 volts. Ponceau Stain In addition to the Western Blot we will be performing, we would like to see the total protein band pattern for each sample. Coomassie staining would ruin the membrane for the Western Blot, so instead we are staining with a soluble dye called Ponceau Red that will be removed during the blocking step. 1. Disassemble the apparatus. 2. Place the blot (labeled side up) in a square Petri dish and just cover (do not fill the dish) with Ponceau red stain. Let sit for 5 minutes. 3. During the staining, discard the gel and two sheets of filter paper and rinse the fiber pads (these are NOT disposable) and the rest of the apparatus unit with plenty of tap water and put away. 4. After the 5 minutes (you will not see bands yet), rinse the blot briefly with distilled water. At this point you will see protein bands. Mark the positions of the molecular weight markers with pencil. You should see a band of 60 kD molecular weight in the positive clones that is not present in the negative control or negative clones. 5. Place the blot back in Petri dish. Blocking The blocking step of the Western Blot is critical to prevent antibody from sticking nonspecifically to the nitrocellulose membrane. 1. Pour the water out of the Petri dish, and add 10 ml blocking solution (TBST + 5% powdered milk). 2. Incubate 15 min on an orbital shaker at 50 rpm. 3. Wash briefly in two changes of TBST. Add fresh TBST, wrap the Petri dish with Parafilm and store at 4˚C until next week.
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12 Expression of Fusion Protein from Positive Clones and Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE) and Immunological Analysis (Western Blot): Part 2 oal: Today we will finish the Western Blot that was started in the last laboratory session.
G I. INTRODUCTION
Probing the Western Blot is very similar to the protocol performed in Lab Session 7C, “Colony Hybridization with a Monoclonal Antibody Probe,” except the protein on the membrane for the Western Blot was derived from the polyacrylamide gel, rather than the colony lift. Since the cellular proteins of each clone were separated by SDS-PAGE, we will be visualizing the specific protein band corresponding to the EGFP::GST fusion protein, rather than just seeing that the protein was made by a specific clone as in the colony lift experiment. For a review on how the GST::EGFP protein is detected on the membrane, revisit Figure 7.5. In brief, following protein transfer to the nitrocellulose membrane, the membrane is blocked with a protein that will not interact with the primary antibody (such as casein found in skim milk or bovine serum albumin). Primary antibody is added and allowed to bind to the specific epitope on the EGFP protein, and unbound antibody is washed away. Secondary antibody (goat anti-mouse) conjugated to horseradish peroxidase (GAMP) is added and allowed to bind to the primary antibody, then excess is washed away. Finally, the colorimetric substrate chloronaphthol is added. Protein samples that contain the GST::EGFP fusion protein will reveal a purple band of approximately 56 kD molecular weight. Clones that did not express EGFP (negative clones) will show no band. Figure 12.1 shows a flowchart of the steps in performing a Western Blot.
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transfer to nitrocellulose membrane
polyacrylamide gel of cellular protein
incubate w/ EGFP-specific mAb. wash unbound mAb.
wash and block
membrane with protein imprint
incubate w/ GAMP. wash.
develop
positive bands “light up”
Fig. 12.1 Western Blot experimental flowchart. Samples are separated by SDS polyacrylamide gel electrophoresis, then transferred to a nitrocellulose membrane. The membrane is washed and blocked, then incubated with the primary antibody, α-EGFP. Excess primary antibody is washed off, and the secondary antibody bound to horseradish peroxidase (goat anti-mouse peroxidase or GAMP) is added. Excess secondary antibody is washed off, then the blot is “stained” using the peroxidase substrate, chloronaphthol.
II. LABORATORY EXERCISES A. SDS-Polyacrylamide Gel Electrophoresis and Western Blot: Part 2 Incubation of the Blot with Antibody Against GFP (Primary Antibody) 1. Obtain an aliquot of the anti-GFP. Store the antibody on ice. 2. Retrieve your blot: It has been incubating at 4˚C in TBS-T since last week. From now on incubations are performed at room temperature. 3. Wash the blot briefly in one change of TBS-T. 4. Make a 1:1000 dilution of the anti-GFP by mixing 7.5 µl antibody with 7.5 ml TBS-T. Mix and add to washed blot. Incubate on an orbital shaker for 1 hr. 5. Wash briefly in two changes of TBS-T. Wash for an additional 10 min with two changes of fresh TBS-T. Incubation with Goat Anti-Mouse Peroxidase (Secondary Antibody) 1. Discard the final wash. 2. Make a 1:500 dilution of the goat anti-mouse antibody conjugated to horse radish peroxidase (GAMP) by mixing 15 µl with 7.5 ml of TBS-T. Add to the blot and incubate for 1 hr on an orbital shaker.
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3. Wash briefly in two changes of TBS-T. Wash for an additional 10 min in two changes of fresh TBS-T. Colorimetric Detection of Peroxidase Activity 1. Add about 10 ml of peroxidase substrate to the tray containing the blot. Color development (dark blue) should be evident within seconds to minutes. 2. Stop the reaction by rinsing with distilled water. 3. To preserve color, dry the blot between two pieces of filter paper. 4. Record the results in your notebook. Positive clones should show a band of approximately 60 kD molecular weight, while negative clones should show no bands. In the positive clones, you may see some smearing or distinct bands below the 60 kD band. This is most likely due to partial degradation of the fusion protein.
B. Replica Plate Positive Clone Replica plate one of your positive clones onto LB/kan so you will have a fresh inoculum for next week. Label, tape shut, and place inverted in the 37˚C incubator. Your instructor will save it in the refrigerator after an overnight incubation.
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13 Extraction of Recombinant Protein from Escherichia coli Using a Glutathione Affinity Column his week you will purify the GST::EGFP fusion protein by affinity chromatography. You will need to inoculate one of your clones into liquid culture at least one day before your lab. You or an instructor will induce protein expression with IPTG 3-4 hours before your regular lab period.
T
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13A Interim Laboratory Session
I. LABORATORY EXERCISE A. Inoculate Cultures for Protein Purification noculate one positive clone (heavily) into 2 ml LB/kan broth in a snap-cap tube. Do this by picking bacterial growth from your master plate with a sterile toothpick and dropping it into the tube containing LB/kan. BE SURE TO LABEL THE TUBE WITH YOUR STATION NUMBER AND LAB DAY. If you are not completely confident that you have a positive clone, you should use pAD1 as your clone for the protein purification. Place the tube in a designated rack. Inoculated cultures will remain refrigerated until the evening before your lab, when your instructor will place them in the 37˚C shaking incubator overnight.
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13B Extraction of Recombinant Protein from Escherichia coli Using a Glutathione Affinity Column oal: Today you will lyse bacterial cells that have been induced with IPTG to express the GST::EGFP fusion protein, and prepare a crude cellular homogenate. You will then use affinity chromatography to purify the GST::GFP fusion peptide encoded by your clone. This is achieved by taking advantage of the affinity of the glutathione-S-transferase (GST) moiety of the fusion protein for the small molecular weight molecule, glutathione.
G I. INTRODUCTION
The first step of purifying a cellular protein is lysing and homogenizing the cells. This will be accomplished by treatment with the enzyme lysozyme, followed by multiple freeze–thaw cycles, followed by sonication. Freezing and thawing bacterial cells lyses them by disrupting the cell membrane, releasing soluble periplasmic and cytoplasmic components. The mechanical action of sonication further disrupts the cells and separates cellular proteins and lipids. It also serves to shear the DNA. Following this homogenization of cellular components, our target protein, the GST::EGFP fusion protein, can be purified by the technique of affinity chromatography. Affinity chromatography is a commonly used method for purifying recombinant proteins. The use of affinity chromatography has become widespread due to the development of numerous expression vectors containing sequences that may be used as affinity tags. These expression vectors encode a gene whose protein may be used as an affinity tag (also called a fusion tag) either directly upstream or downstream of the multiple cloning site. The gene of interest must be inserted in the correct orientation and reading frame with respect to the affinity tag. Once protein expression is induced, a fusion protein will be produced. The fusion protein will contain two polypeptide moieties; one corresponding to the affinity tag, and one corresponding to the protein of interest. Numerous affinity tags exist. Some of the most common include glutathione-S-transferase (the tag we use), hexahistidine (a series of 6 histidine residues), and a cellulose binding domain. pET-41a(+) contains the gene for glutathione-S-transferase (gst) directly upstream of the polylinker region, and after a gene of interest (in our case, egfp) is inserted into the polylinker in the correct orientation and reading frame, a fusion protein can be expressed. The fusion protein we will express is GST::EGFP. We will take advantage of the GST portion of the GST::EGFP fusion protein to purify it via affinity chromatography.
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The principle of affinity chromatography relies on the binding of a biospecific ligand to the molecule of interest. The ligand for GST is glutathione, which is covalently attached to a support matrix to allow separation of glutathione binding molecules from other cellular components. The principle steps of affinity chromatography are described below, and are illustrated in Figure 13.1. The specific details of our purification are indicated in italics. 1. Injection of sample. The sample contains a mixture of proteins and other cellular components along with the recombinant protein of interest. The sample you will use is a crude homogenate of the induced culture containing your recombinant DNA. 2. Adsorption of molecules with affinity for the ligand. Only the GST::EGFP fusion protein will bind to the glutathione linked affinity matrix. 3. Wash impurities from the column. Molecules with no affinity for the ligand are washed from the column. Cellular molecules other than the GST::EGFP fusion protein will flow through the column, and the GST::EGFP fusion protein will remain bound to glutathione in the column. 4. Elution of the target molecule(s) from the column. The GST::EGFP fusion protein will be eluted from the column using a solution of reduced glutathione.
A
B
C
D
Fig. 13.1 The principle steps of affinity chromatography. A. Sample injection. B. Adsorption of target molecule(s). C. Washing of impurities. D. Elution of target molecule(s).
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The affinity column will be packed with a slurry we will refer to as the “affinity resin,” illustrated in Figure 13.2. The substance that makes up the affinity resin has 3 covalently bound components; the ligand, the matrix, and the spacer.
matrix
spacer
ligand
Fig. 13.2 The affinity resin is composed of a biospecific ligand, a support matrix, and a biochemically inert spacer.
The biospecific ligand of the affinity resin specifically binds the molecule of interest. Depending on the type of molecule to be purified, various classes of ligands may be used. For example, if an enzyme is to be purified, the ligand in the affinity phase may be a substrate, inhibitor, or cofactor of the enzyme. If an antibody is to be purified, the cognate antigen may be used as the ligand. If a nucleic acid is to be purified, ligands could be complementary sequence or, possibly, DNA binding proteins. Affinity resin can be made in the laboratory by chemical cross-linking of reactive resins with ligand, but you will use a commercially available affinity matrix. Because your protein is tagged with the enzyme glutathione-S-transferase, the biospecific ligand you will use is the substrate, glutathione. The support matrix functions to suspend the ligand in the column. The matrix must be rigid, biochemically inert, and have a high surface to volume ratio. The most commonly used support matrix is sepharose. Sepharose is composed of small agarose beads and is available in various sizes. The spacer is a carbon chain that links the ligand to the matrix. The purpose of the spacer is to present the ligand to the molecule of interest. If no spacer was present, or if a spacer was too short, the ligand would be embedded in the matrix and might not be available for binding to the molecule of interest. Conversely, a spacer that is too long could interact with or have affinity for undesired molecules. The ideal spacer would be just long enough to present the ligand, but short enough to remain biochemically inert. A final consideration when designing an affinity purification protocol in your own research laboratory is the issue of column capacity. Column capacity refers to the number of ligand molecules available for binding. In practice, that would mean the amount of affinity resin packed into the column. If there is not enough ligand for all of the target molecules in the sample to bind, then some of the target molecules will flow through the column and be lost. Conversely, if there is an excessive amount of ligand binding sites compared to target molecules in the sample, non-target sample molecules with a lower affinity for the ligand may be able to bind. If these non-target molecules are able to remain bound during the wash steps, they will be eluted with the sample. This, of course, could lead to impurities in the purified product. Optimally, the number of ligand binding sites would be exactly equal to the number of target molecules in the sample. In practice, this would be impossible to achieve. Column capacity needs to be optimized empirically in most cases because expression levels of various proteins expressed from various promoters under varying growth conditions differ. The diagnostic SDS-PAGE will help you to analyze the purity of your sample. If the eluate does not appear to be pure, or if you lost a great deal of target protein in the washes, then you should modify your purification protocol either by adjusting column capacity or by adjusting the stringency of the washes.
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II. LABORATORY EXERCISES A. Growing Bacterial Suspension Cultures for Fusion Protein Purification Your instructor will inoculate your overnight culture into 100 ml of 2× YT/kan/IPTG medium, and incubate at 37˚C, 3–4 h.
B. Harvesting IPTG-Induced Cultures 1. Divide your culture between two Oak Ridge tubes (i.e., 35–45 ml in each tube). Excess culture can be discarded in the “bacterial graveyard.” 2. Pellet the bacteria by spinning for 5 min at 10,000× g in a high-speed centrifuge at 4˚C. 3. Pour off the clear supernatant and discard. 4. Transfer the two pellets to a single microcentrifuge tube using the following method: Add about 0.75 ml of 2× YT medium (or LB) to one bacterial pellet. Pipette and expel to resuspend the pellet. Transfer this to the second pellet and pipette and expel to resuspend. Now transfer the thick soup to a 1.5-ml microcentrifuge tube. Repeat this procedure with 0.5 ml of 2× YT to transfer any remaining bacteria. 5. Balance the weight of the bacteria-containing tube with that of another (water-containing) tube and spin for 1 min in a microcentrifuge at 13,000 rpm to pellet the bacteria. Pour off the clear supernatant. Invert the bacteria-containing tube over a paper towel and leave for about 5 min to remove residual liquid.
C. Breaking Open Bacterial Cells 1. Add 0.75 ml of ice-cold 1× GST Bind/Wash Buffer (with Pefabloc, a protease inhibitor) to the pellet. Vortex to suspend the pellet. 2. Add 15 µl of lysozyme (10 mg/ml). Vortex and incubate on “wet” ice (H2O) for 15 min. 3. Place the microcentrifuge tube on a bed of “dry” ice (CO2) to freeze. 4. Thaw the tube by warming it in the palm of your hand. As soon as the sample melts, freeze it again on dry ice. 5. Thaw the tube as described above in step 4, and store it in liquid form on wet ice. Freezing and thawing helps release the fusion protein by disrupting the bacterial cell wall. CAUTION Put on protective ear guards before activating the sonicator in the next step. 6. You will use a sonicator with an immersible tip (probe). Note: There may be a maximum power setting for this tip; make sure you do not exceed this setting. The conditions are not specified here because they vary depending on the particular sonicator used—most likely you will use a low setting because of the small volume being sonicated. Rinse the tip with distilled water. Immerse the tip into the bacterial solution, but do not let the tip come into contact with the sides of the tube. Sonicate the ice-cold solution for 15 sec and return the tube to the ice. Sonication will heat up the solution and it is important to sonicate for short periods of time. Repeat this procedure three times, returning the tube to ice between sonications. This fraction represents the crude homogenate (CH). Sonication helps to release the fusion peptide from the bacterial cells and reduces the viscosity of the solution by shearing chromosomal DNA.
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7. Record the approximate volume of the crude homogenate: _____. 8. Take a sample of crude homogenate for the Bradford protein assay (10 µl). Label the tube and freeze at −20˚C. 9. Remove an additional 10 µl of the crude homogenate for analysis by SDSPAGE. Add 10 µl of 2× sample buffer (containing bromophenol blue). Label the tube and freeze at −20˚C.
D. Removing Insoluble Debris from the Crude Homogenate 1. Spin the crude homogenate at 13,000 rpm in a refrigerated microcentrifuge or in a microcentrifuge kept in a cold box (4˚C) for 15 min to remove insoluble debris. The insoluble debris represents cells that were not thoroughly sonicated. If the pellet is very large, resuspend the pellet, sonicate for additional time, and repeat the centrifugation step. 2. Transfer all of the supernatant to a fresh microcentrifuge tube and store on ice.
E. Purifying Protein by Affinity Chromatography The resin (affinity matrix) and buffers should be allowed to equilibrate to room temperature before use. 1. Your instructor will provide you with a slurry of the GST-Bind Resin (affinity matrix) in a sealed 15 ml test tube. You should have approximately 2 ml of settled bead volume. Note: Be gentle—the beads are fragile. 2. Assemble the chromatography column. Attach the stopcock to the bottom of the column and the reservoir collar to the top. Mount the assembled column on a ring stand. 3. Make sure the stopcock is closed. Invert the GST-Bind resin to gently mix. Add the uniform slurry of affinity matrix material to the column. Open the stopcock to allow the buffer to drip through; do not allow the buffer to drop below the top of the settling beads. Close the stopcock once the buffer level is just above the affinity resin. Also, make sure your column does not leak. If it does, you may have to replace the stopcock. 4. Wash the column with an additional 10 ml of GST Bind/Wash Buffer. Keeping the stopcock wide open, allow the liquid level to drop until just above the bed of resin. Close the stopcock. 5. Add 3.25 ml of room temperature GST Bind/Wash Buffer to your thawed sample. 6. Add your sample to the column and open the stopcock. Collect the flowthrough fraction in a test tube labeled “FT” and store on ice. Close the stopcock once the buffer level is just above the settled resin. In affinity chromatography, the binding of the fusion protein to the ligand requires a slow flow rate. Allowing the sample to flow through the column by gravity flow achieves this. Forcing the sample through the column at a more rapid rate will decrease the binding efficiency of the target molecule to the ligand. 7. Add 10 ml GST Bind/Wash Buffer to the column, open the stopcock and collect the wash fractions, 1 ml per tube labeled W1-W10 and freeze. Most of the Escherichia coli proteins will wash through in this step. Close the stopcock once the buffer level is just above the settled beads. 8. Wash the column with an additional 10 ml of GST Bind/Wash Buffer to remove any non-GST proteins left behind. Close the stopcock once the buffer level is just above the settled beads. Discard the flow-through solution.
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9. Label 12 microcentrifuge tubes for the eluted samples (eluates) as E1–E12. 10. Add 6 ml GST elution buffer to the column. Open the stopcock and collect ~0.5 ml fractions in your labeled tubes. 11. Place the tubes with your washes and eluates on the UV transilluminator in a dimly lit room. Be sure to wear a UV-protective face shield. Record which fractions have green fluorescence in your notebook.
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14 Analysis of Purification Fractions oal: Today you will analyze your purification fractions by SDS-PAGE and quantify protein concentrations using the Bradford assay. SDS-PAGE will allow you to estimate purity of the eluted fractions and to determine whether any of the recombinant fusion protein was lost during the wash steps. The Bradford assay will be used to determine how much fusion protein was recovered.
G I. INTRODUCTION
In Lab Session 11 you performed an SDS-PAGE on extracts from your transformants and then transferred the protein to a nitrocellulose membrane and probed with α-EGFP to confirm the presence of the GST::EGFP fusion protein. In that particular case, we were not concerned with the presence or absence of any other protein(s) in the sample—we simply wanted to confirm that GST::EGFP was expressed. The purpose of the SDS-PAGE we will perform in this laboratory session is to determine which washes, if any, contain fusion protein and to determine whether the fusion protein is free of contaminating protein in the eluted fractions. We will not use antibodies because we need to visualize all of the proteins in the wash and eluted fractions. Figure 14.1 (next page) shows an example of a gel stained for total protein. The crude homogenate includes all cellular proteins, including the fusion protein. Ideally the washes will contain all cellular proteins EXCEPT the fusion protein, and the eluate fractions will contain ONLY the fusion protein. Presence of excess fusion protein in wash fractions indicates that the column did not have enough affinity resin or that the flow-through rate was too fast for the fusion protein to bind to the resin. Presence of additional protein bands in the eluate fractions can either indicate that the column was not washed completely, or that there was proteolysis of the fusion protein. How might a researcher distinguish between additional bands arising from contaminating protein versus bands arising from proteolysis? Column capacity conditions could be optimized to improve the yield of pure fusion protein, based on the results of the SDS-PAGE. Protein concentration assays measure the concentration of total protein in a sample. A number of assays exist for protein concentration. For each of these assays, protein concentration is estimated by comparing the value of an unknown sample to a standard curve created by assaying a series of known concentrations of a reference protein such as BSA (bovine serum albumin). The most commonly used assays are the Bradford assay and the BCA assay. Each assay has its own strengths and weaknesses.
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2
3
4
5
6
7
8
9
10
Fig. 14.1 SDS-PAGE of EGFP::GST purification fractions (10% polyacrylamide). Lane 1, NEB molecular weight marker premixed with sample buffer; lane 2, crude homogenate; lane 3, wash #1; lane 4, wash #2; lane 5, wash #3; lane 6, eluate 3; lane 7, eluate 4; lane 8, eluate 5; lane 9, eluate 6; lane 10, eluate 7.
The Bradford assay is the one we will use to determine the protein concentrations. This assay uses Coomassie G-250, which binds primarily to basic (especially arginine) and aromatic amino acid residues. When protein is present in an acidic solution, the color of bound Coomassie G-250 shifts from brown to blue. The color shift is proportional to the amount of protein in solution, and can be measured spectrophotometrically. This assay is able to detect and quantify protein concentrations on the order of 1–1400 µg/ml. The disadvantage of this assay is that it is incompatible with most detergents and some common protein buffers. If our protein purification method required the use of detergents, the Bradford assay would not be a good choice. If no detergent is used in the purification method, and a compatible buffer is used, it is an excellent method. The BCA (bicinchoninic acid) assay has the advantage that it is compatible with various detergents and buffers. However, this assay is is based on interactions with tyrosine residues in the protein sample. For a mixed sample of protein, this is useful, but for individual proteins this can be problematic. A protein sample that has a below-average amount of tyrosine residues will give an artificially low reading, and a protein sample that has an above-average amount of tyrosine residues will give an artificially high reading. For this reason, the BCA assay is not advised for measuring the concentration of a single protein, especially if the tyrosine content is not known. This assay has a sensitivity range of 20–2000 µg/ml. The classic Lowry assay is similar to the BCA assay and is also dependent on the presence of tyrosine residues. The Lowry assay is not compatible with detergents and a number of commonly used buffers. It is not generally used because it has the negative attributes of both the Bradford and BCA assays without any additional advantage. We are using the Bradford assay because we are measuring the concentration of a single, purified protein (the GST::EGFP fusion protein), therefore the BCA
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assay would not be ideal. Because the fusion protein is a soluble protein, detergents are not necessary for its isolation, and thus the Bradford assay is compatible with the protein sample.
II. LABORATORY EXERCISES A. SDS-PAGE of Purified Fusion Protein SDS-PAGE will provide a qualitative analysis of sample purity. Retrieve your samples that you took last week from the −20˚ freezer. Label microcentrifuge tubes and aliquot the following: 1. 2. 3. 4. 5. 6. 7. 8. 9. 10.
Molecular weight marker premixed with sample buffer (10 µl) Crude homogenate (20 µl—already has sample buffer) Wash #1 (10 µl sample +10 µl sample buffer)* Wash #2 (10 µl sample +10 µl sample buffer)* Wash #3 (10 µl sample +10 µl sample buffer)* Eluate 3 (10 µl sample +10 µl sample buffer)∧ Eluate 4 (10 µl sample +10 µl sample buffer)∧ Eluate 5 (10 µl sample +10 µl sample buffer)∧ Eluate 6 (10 µl sample +10 µl sample buffer)∧ Eluate 7 (10 µl sample +10 µl sample buffer)∧
Once the aliquots have been made, heat the samples for 10 minutes at 95˚C or higher before loading. Use tube tabs to lock caps. (The molecular weight marker has already been heated and only needs to be heated 1 minute.) This will help to prevent aggregation. Run the gel at constant current until the dye front is at the bottom of the gel (as described in Lab Session 11). Staining the Gel CAUTION: Wear gloves and lab coat to prevent blue staining. 1. Detach the gel from the upper reservoir. Be careful not to squeeze the gel. 2. Pull apart the two pieces of plastic gently (as described in Lab Session 11). The gel should stick to one plate. 3. Cradle the plate in your hand with the gel side facing up. While holding it over your staining tray, direct a stream of distilled water (from a squirt bottle) under the gel, to loosen it. Tilt the plate to allow the gel to drop into the staining tray (a square Petri dish or plastic top of a pipette tip box makes a convenient tray). 4. Place the gel in the tray and rinse 3 times (5 minutes each) with dH2O with gentle shaking. Label the tray with your station number and lab day on a piece of tape. 5. Mix GelCode Blue Reagent by inverting gently immediately before using. 6. Add approximately 20 ml of stain reagent (enough to cover gel, but do NOT fill the tray with it!) and shake tray for an hour. 7. Replace the stain reagent with dH2O, and continue shaking. You can probably see the bands at this point, but this step enhances stain sensitivity and weak protein bands will continue to develop. The water may need to be changed several times for optimal results. The gel can be left in water for as long as necessary—15 minutes to overnight. 8. Photograph your gel and analyze the results.
*
If any of your washes appeared to have green fluorescence, run those instead. If eluate fractions other than these had green fluorescence, run those instead.
∧
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B. Bradford Protein Concentration Determination Assay of Purification Fractions Start this assay while your SDS-PAGE is running 1. Prepare dilutions of BSA for standard curve as described below: You will be given a tube with 100 µl of a 1-mg/ml BSA solution. Prepare a dilution series of BSA standards in 5 tubes (labeled 1–5) according to the scheme in Table 14.1. Attach a new pipet tip to the micropipet each time to make the dilutions. Be sure to mix each tube before transferring a portion to the next tube.
TABLE 14.1 BSA Dilutions Tube 1 2 3 4 5
Dilution
Protein concentration
100 µl of stock (1 mg/ml) + 300 µl dH20 150 µl from tube 1 + 150 µl dH20 150 µl from tube 2 + 150 µl dH20 150 µl from tube 3+ 150 µl dH20 150 µl from tube 4 + 150 µl dH20
250 µg/ml 125 µg/ml 63 µg/ml 31 µg/ml 16 µg/ml
2. Prepare samples IN TRIPLICATE as shown in Table 14.2.
TABLE 14.2 Composition of Assay Wells for Bradford Analysis Sample Crude homogenate
Sample Volume
Vol. of Water Added to Dilute Sample (d1)
Vol. of Sample & Water Added to Well (d2)
10 µl
90 µl
10 µl diluted sample + 30 µl H2O
Wash Fractions Wash1 Wash 2 Wash 3
1 ml 1 ml 1 ml
40 µl undiluted sample 40 µl undiluted sample 40 µl undiluted sample
Eluate Fractions
Eluate 3 Eluate 4 Eluate 5 Eluate 6 Eluate 7
500 µl 500 µl 500 µl 500 µl 500 µl
-
10 µl undiluted sample + 30 µl H2O 10 µl undiluted sample + 30 µl H2O 10 µl undiluted sample + 30 µl H2O 10 µl undiluted sample + 30 µl H2O 10 µl undiluted sample + 30 µl H2O
3. Add 40 µl of dH2O in well A1, B1, and C1; 40 µl of BSA dilution 5 in well A2, B2, and C2; 40 µl of BSA dilution 4 in well A3, B3, and C3; and so on for the standards (from most dilute to most concentrated). 4. Immediately add triplicate samples to rows D, E, and F (40 µl). Follow the scheme shown in Table 14.3. If you noted last week that eluates other than E3-E7 fluoresced green, use those eluates instead.
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Lab Session 14 TABLE 14.3 Schematic of Microtiter Plate
A B C D E F G H
1
2
3
4
5
6
7
8
9
Water Water Water CH CH CH
BSA5 BSA5 BSA5 W1 W1 W1
BSA4 BSA4 BSA4 W2 W2 W2
BSA3 BSA3 BSA3 W3 W3 W3
BSA2 BSA2 BSA2 E3 E3 E3
BSA1 BSA1 BSA1 E4 E4 E4
E5 E5 E5
E6 E6 E6
E7 E7 E7
10
11
12
5. Add 160 µl Bradford reagent to each water, BSA and sample well. Use a multichannel pipettor* if available. Be sure not to form bubbles. Incubate 10 minutes at room temperature, and read the plate at 595 nm. It is important that the reactions for the standard curve be read at the same time as the samples because the Coomassie G-250 can continue to react with protein, changing the absorbance. 6. Calculate the protein concentration and total protein amount of each sample. Calculate the Concentration and Amount of Protein in Each Sample 1. Create a standard curve. Graph a standard curve derived from the blank and BSA dilutions. Graph the known concentrations on the X axis and the A595 on the Y axis. 2. Determine the protein concentration in each assay well (Ca). Take the A595 average of each triplicate assay reading. If one of the numbers is far off, and you suspect that you made a mistake in pipetting, you may discard that reading. Plot the average absorbances on the standard curve and interpolate the unknowns to determine their concentrations. Alternatively, use a computer program such as Microsoft Excel to calculate the slope of the standard curve m = ∆Y/∆X or slope = ∆A595/∆concentration. Then plug in y = mx + b or A595 = slope ¥ concentration (b = 0 in this case). The concentration you calculate here is the concentration of the sample in the assay well (Ca). Recall that you diluted the crude homogenate and the eluates for the assay, so to determine the protein concentration in the original sample (Co), further calculations are necessary. 3. Determine the concentration in the original sample (Co). Co(µg/ml) =
Ca(µg/ml) d1d2
Where Co is the protein concentration in the original sample (µg/ml), Ca is the protein concentration in the assay microtiter well, and d1 and d2 are the dilution factors referred to in Table 14.2. d1 is the dilution of original sample, which equals the sample volume/(sample volume + distilled H2O dilution volume), and d2 is the dilution of sample in the assay tube, which equals the volume of sample added to the assay tube/total volume added to the assay tube. For example, for the crude homogenate sample, d1 = 10/(10 + 90) = 0.1 and d2 = 10/(10 + 30) = 0.25. The wash fractions were not diluted, and the eluates only have a d2.
*
Multichannel pipettors are notorious for dropping the pipet tips. Be sure to handcheck that each tip is on tightly.
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4. Calculate total protein. Total protein = protein concentration × total volume You recorded the total volume of the crude homogenate in Lab Session 13.C.7.
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APPENDIX
1 Equipment
SHARED EQUIPMENT ● ●
● ● ● ●
● ● ● ●
● ● ● ● ●
● ● ● ● ● ● ● ●
−80˚C degree freezer −20˚C degree freezer (if the freezer has an automatic defroster, store all enzymes in insulated coolers, such as Stratacoolers, inside the freezer) Refrigerator Centrifuge UV crosslinker (Stratalinker) Floor model shaker incubator or table top shaker incubators for 100-ml bacterial cultures 37˚C incubator for bacterial plates and enzymatic reactions Microwave oven for melting agarose Sonicator with micro-immersion tip (such as the Branson Sonifier® Model 250) GeneQuant (or other spectrophotometer that measures absorbance at 260 and 280 nm) with quartz cuvette Orbital benchtop shaker Microtiterplate reader (optional) Spectrophotometers (if no microtiter plate reader available) Hybridization oven (or water bath with sealable baggies) UV transilluminator with gel documentation system (or UV transilluminator in darkroom with camera) PCR thermocycler Autoclave Speed Vac (optional, for evaporating ethanol from DNA preparations) Large water baths for cooling media to 60˚C PH meter Top-loading balance Ice machine Water purification system (such as Dracor or Millipore)
AT EACH LAB STATION Each lab station should have the items detailed in the Station Checklist in Lab Session 1.
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2 Prep List
NOTES TO PREP STAFF Plasmids and E. coli Host Strains he three plasmids you will need are pET-41a(+) (Novagen cat. # 70556-3), pEGFP-N1 (Clontech cat. # 632318), and pAD1 (request from Dr. Sue Carson at
[email protected]; include “pAD1 request” in subject heading). The two host strains used are the expression host BL21(DE3) (Novagen cat. # 70235-3) and the non-expression host NovaBlue (Novagen cat # 70181-3). The catalog numbers given are for chemically competent cells. If you choose to make the competent cells, you may purchase regular, non-competent cells. Alternatively, other similar host strains may be purchased from other companies. Keep in mind that you will need one strain for isolating DNA (such as NovaBlue), and that strain should not be specialized for protein expression. You will need a second strain that allows transcription from the T7 promoter: this means that the expression strain must be a λ lysogen (such as BL21(DE3)). Before the course begins, you will need a freezer stock of the following E. coli host strains (or similar strains) carrying the following plasmids.
T
E. coli Host Strain NovaBlue NovaBlue BL21(DE3) BL21(DE3)
Plasmid pEGFP-N1 pET-41a(+) pET-41a(+) pAD1
Purpose plasmid isolation plasmid isolation negative control for fusion protein expression positive control for fusion protein expression
Antibiotics Each of the plasmids use kanamycin for selection. Make a stock solution of kanamycin at 50 mg/ml in water, filter sterilize, and store at –20. Add kanamycin to media at a dilution of 1:1000 from the stock solution (i.e., 1 ml in 1 liter) after autoclaving. It is critical to wait for media to cool to approximately 60˚C before adding antibiotic. Adding antibiotic while the medium is too hot can cause inactivation of the antibiotic. It is also important to make sure that the antibiotic being used is effective; bad lots do occur. This can be tested by inoculating a freshly made LB/kan plate with wild type E. coli (lacking a plasmid containing antibiotic resistance).
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Aliquots Restriction enzymes and buffers should be aliquoted for individual use. Whenever feasible, make more aliquots than are needed for class. Students often misplace aliquots or need to repeat a procedure. Extra aliquots come in handy. Plan on 30% extra aliquots for Lab Sessions 1–4, especially if students do not have a great deal of prior experience with micropipettors. Individual aliquots should be labeled. Either label by hand, or use laser-printed Tough-TagsTM.
Bacterial Waste Solid Solid bacterial waste includes tubes, tips, plates, etc., that have been used to grow bacteria, or that have come in contact with any living bacteria/biohazardous waste. Recombinant DNA should also be considered as biohazardous. The prep staff should provide several white plastic buckets (approximately 25-gal size) and/or large aluminum wire baskets. Line buckets/baskets with appropriate-sized orange BIOHAZARD bags (Fisher, Pittsburgh, PA). Replace the bags when a little over half full. Do not allow students to place long disposable pipettes in bags—the bags will be punctured and you will have to rebag the bacterial waste. Pipettes can go in a bucket filled with dilute Clorox. You can then dispose of them in the regular trash. All BIOHAZARD bags must be autoclaved before disposing. Note: Do not allow regular trash to be placed in orange BIOHAZARD bags. It can be dangerous to autoclave and puts an additional burden on the prep staff. For convenience, students should have a small bench-top biohazard container lined with an orange autoclave bag at each station. This should be used for pipet tips and other small items that are used at the lab bench. Students should empty their small biohazard waste containers into the large common container at the end of each lab. Liquid Prepare a carboy labeled “bacterial graveyard” for liquid bacterial waste only. Treat with Clorox (approximately 15–20% by vol). Wait 10–20 min. Bacteria are dead if the liquid has cleared and cell debris has fallen to bottom of the container. Decontaminated liquid may be poured down the drain.
Autoclaving Autoclaved materials should be ready the day before the material is needed. Liquid Fill bottles only four/fifths full. Caps should be loose to prevent pressure build-up and bursting. Tighten the caps fully only after the solution has cooled to room temperature. Liquids should be autoclaved for 45–55 min, depending on the volume. Antibiotics should be added to media after autoclaving. Agar will settle following autoclaving. Media should be gently swirled and allowed to stand 5–10 min before pouring into plates to allow bubbles to subside. Media should be cooled to 60–65˚C before pouring, and can be stored in a water bath at this temperature. Label plates with three stripes for LB/kan and one stripe for LB, no antibiotics. Biohazard Waste Orange autoclave bags must be sealed and labeled. If leakage is a problem, they may have to be double bagged or absorbent material added (e.g., vermiculite, bentonite, etc.).
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Dry Goods All pipet tips and microfuge tubes should be autoclaved before use. Students may rack their own pipette tips during times they are waiting to use a piece of equipment. They will leave items labeled with their initials on autoclave indicator tape to be autoclaved in designated autoclave trays. These items should be autoclaved so that they are ready for the next lab.
General Lab Preparation Designate an area or cart for reagents for student use. Group together materials needed for a particular lab session. Use 5 × 7 note cards to label all reagents and include information such as “for lab 7, part 3a.” Include phrases such as “Return when finished,” “Do not throw away, label with your group number and store at 4˚C,” or “Discard following use.” Be sure to also label each individual aliquot, as students will often mix up tubes! Clear directions provided whenever possible will help avoid problems and are especially important to ensure a safe working environment. Periodically check the orange BIOHAZARD bags and remove them for autoclaving when they are half full. Make sure that deionized water carboys are full.
Students Students will ask for help and for more reagents, particularly during the first 4 sessions. Students should be advised not to go to the prep room. Advanced preparation and having additional aliquots on hand will help to avoid frustration.
SUPPLIES AND REAGENTS Supplies and reagents for each week are listed in this appendix according to lab session. There are certain supplies and reagents that should be available at all times. These will not necessarily be listed under each lab session, but should be available for students and prep staff every lab session. These are listed below.
Supplies and Reagents for General Use ●
● ● ● ● ● ●
● ●
●
● ● ● ●
Latex gloves (small, medium, large, and extra-large); powder-free gloves are preferable for working with nylon and nitrocellulose blots Pipet tips (all sizes) Microcentrifuge tubes Sterile distilled/deionized water (carboys) 95% ethanol (stored at –20˚C) 70% ethanol (carboy) Agarose (can be kept at general-use area containing top-loading balance and microwave) Petri dishes (square and round) Kanamycin (kan) (50 mg/ml stock solution in dH2O, this is a 1000× solution), filter-sterilized, stored at –20˚ C Isopropyl-ß-D-thiogalactopyranoside (IPTG) (20 mg/ml stock solution in dH2O), filter-sterilized, stored at –20˚ C 50 ml conical tubes 15 ml conical tubes Oak Ridge tubes or equivalent for centrifugation at 15,000× g. Ice buckets
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● ● ● ● ●
Plate spreaders or glass rods for making plate spreaders (1 per student) sterile toothpicks and/or wire loops for inoculating cultures LB broth (on hand in prep room) LB/kan agar plates (stored at 4˚C) TE buffer (on hand in prep room) 5× TBE (carboy) Ethidium bromide (1 mg/ml) (store protected from light in an amber container or wrapped in foil)
Recipes for General Use LB Broth 1. Mix the following: Compound
Amount per liter
Tryptone NaCl Yeast extract Deionized water
10 g 10 g 5g 1000 ml
2. Bring to pH 7.0 with 5N NaOH (approximately 0.2 ml) 3. Autoclave 4. Cool and add antibiotics and other (filter-sterilized) additives, such as IPTG, where necessary To prepare LB agar, follow steps 1 and 2 above, then add 15 g/L agar and swirl. Autoclave, then place in a 60˚C water bath. When the temperature has equilibrated, add antibiotics and other (filter-sterilized) additives where necessary, and pour approximately 20 ml per Petri dish. TE Buffer, pH 8.0 (Tris EDTA) Each solution must be made separately then mixed from stock solutions to make TE buffer. Component
Final Concentration
Amount to add for 1 liter
10 mM 1 mM
10 ml 2 ml
Tris-Cl (1 M, pH 8.0) EDTA(0.5 M, pH 8.0) Deionized water
to 1 liter
To prepare stock solutions: Tris-Cl (1 M, pH 8.0) ● ● ● ●
60.5 g Tris base 450 ml deionized water Bring to pH 8.0 with concentrated HCl Add deionized water to 500 ml
Ethylenediaminetetraacetic acid (EDTA) (0.5 M, pH 8.0) Mix: ● ● ●
● ●
93.05 g of EDTA 450 ml deionized water Bring the pH to 8.0 with NaOH pellets (approximately 20 g). It will take a while to go into solution. Add deionized water to 500 ml Autoclave or filter-sterilize
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5X TBE Stock (Tris-borate-EDTA) Reagent
Amount
Tris Boric acid EDTA dH2O
54 g 27.5 g 4.15 g (or 20 ml 0.5 M EDTA, pH 8) to 1 liter
Place in carboy for students to prepare 1× stocks for their own use. Prepare 6 liters for 20 groups. Ethidium Bromide Stock (1 mg/ml) 1 ml per group, in tinted microcentrifuge tubes, or wrapped with aluminum foil since EtBr is light-sensitive. Students should save the ethidium bromide at their lab station to use in multiple laboratory exercises.
LAB SESSION 1 The students will use the following supplies: ● ● ● ● ● ● ● ●
Sterile distilled/deionized water (at least 1 ml per student) Bovine serum albumin 1 mg/ml (15 µl per student) Nitrocellulose membrane approximately, 2” × 3” (1 sheet per group) Whatman 3MM paper, approximately 2” × 3” (1 sheet per group) Square Petri dish, tip box top, or other convenient tray for staining (1 per group) Waste containers with funnels: for used amido black and destaining solution Amido Black Stain Destaining solution
Recipes Amido Black Stain: The following recipe provides one bottle of stain for general use. Stain may be reused if stored in an airtight container. 1. Combine the following:
Naphthol blue black 10B (Sigma-Aldrich) Methanol Acetic Acid Deionized water
Amount
Final Concentration
0.5 g
0.1%
225 ml 35 ml to 1 liter
45% 7%
2. Filter through fluted filter paper 3. Store at room temperature Destaining Solution: The following provides two bottles for general use; mix and store at room temperature.
Methanol Acetic acid Deionized water
Amount
Final Concentration
700 ml 70 ml to 1 liter
70% 7%
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LAB SESSION 2 Note to prep staff: 1. Two days before lab, streak LB/kan plates of the cloning strain of E. coli (such as NovaBlue) containing the pET-41a(+) and pEGFP-N1 plasmids. a. source of vector DNA: pET-41a(+) in E. coli strain NovaBlue or other appropriate cloning strain b. source of insert DNA: pEGFP-N1 in E. coli strain NovaBlue or other appropriate cloning strain 2. The evening before the lab, inoculate several 100 ml cultures of each strain in LB/kan broth and shake at 37˚ C. Each group will need one 100 ml culture. You may inoculate the strain containing pET-41a(+) for half of the class and the strain containing pEGFP-N1 for the other half of the class. Have available for pre-lab setup: ● LB broth containing 50 µg/ml kanamycin ● LB agar containing 50 µg/ml kanamycin ● E. coli strain NovaBlue (or other K12 strain) containing pET-41a(+) ● E. coli strain NovaBlue (or other K12 strain) containing pEGFP-N1 The students will use the following supplies and equipment: Oak Ridge centrifuge tubes Isopropanol, room temperature ● 70% Ethanol, room temperature ● TE buffer, pH 8.0 (students should save this in their freezer boxes for future labs) ● QIAGEN® QIAfilter Plasmid Maxi Kit (catalog number 12263) The following supplies and reagents are included in the QIAGEN® QIAfilter Plasmid Maxi Kit (catalog number 12263). Each kit contains columns, filters, and reagents for 25 plasmid preps. Buffer solutions are included in the kit, but can also be prepared according to the manufacturer’s instructions. Contents of QIAGEN buffers are listed below with the manufacturer’s permission. QIAfilter Maxi Cartridge (1 per group) QIAGEN-tip 500 (1 per group) ● ●
Buffer
Composition
Storage
Buffer P1 (resuspension buffer)
50 mM Tris-Cl, pH 8.0; 10 mM EDTA; 100 µg/ml RNase A
2–8˚C
Buffer P2 (lysis buffer)
200 mM NaOH, 1% SDS (w/v)
Buffer P3 (neutralization buffer)
3.0 M potassium acetate, pH 5.0
15–25˚C or 2–8˚C
Buffer QBT (equilibration buffer)
750 mM NaCl; 50 mM MOPS, pH 7.0; 15% isopropanol (v/v); 0.12% Triton X-100 (v/v)
15–25˚C
Buffer QC (wash buffer)
1.0 M NaCl; 50 mM MOPS, pH 7.0; 15% isopropanol (v/v)
15–25˚C
Buffer QF (elution buffer)
1.25 M NaCl; 50 mM Tris-Cl, pH 8.5; 15% isopropanol (v/v)
15–25˚C
15–25˚C
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LAB SESSION 3 Note to prep staff: Prior to this lab, you should have collected the pET-41a(+) isolated from students in Lab Session 2, then run each sample on a gel to quality control the plasmid DNA. Pool acceptable samples, quantify, and aliquot to hand back to the students for this lab exercise. The students will use the following supplies: ● ● ● ● ●
● ● ● ●
● ● ●
2.5 µg pET-41a(+) DNA 2.5 µl Nco I enzyme 2.5 µl Not I enzyme 5 µl 10X restriction enzyme buffer 1 ml ethidium bromide stock (1 mg/ml) (see recipe in “Supplies and reagents for general use”) Agarose 2 µl BSA (10 mg/ml) 10X DNA loading buffer (students should save for future labs) 1 kb DNA ladder (Invitrogen, catalog number 15615-024) (students should save for future labs) 5X TBE buffer (see recipe in “Supplies and reagents for general use”) QIAGEN® QIAquick Kit Sterile deionized water
The following supplies and reagents are included in the QIAGEN® QIAquick Kit (catalog number 28104). Each kit contains columns and reagents for 50 DNA “clean-ups.” Aliquot the following amounts for student use: ● ● ● ● ●
195 µl Buffer PB 0.75 ml Buffer PE 55 µl Buffer EB 1 QIAquick column 1 2 ml collection tube
Recipes 10X DNA Loading Buffer Reagent 25% Glycerol 0.1 M EDTA 0.25% Bromophenol Blue
Amount Needed for 50 ml 12.5 ml 10 ml of 0.5 M EDTA 0.125 g
Aliquot 1 ml per tube, 1 tube per group. Students should save loading buffer for future labs. 1 Kb DNA Ladder (Invitrogen, Catalog Number 15615-024) Dilute 1 part DNA ladder with 1 part 10× DNA loading buffer, and 8 parts sterile deionized water. Aliquot 100 µl per group. Students should save extra 1 kb ladder in their freezer boxes for future labs. If an alternative molecular weight ladder is used, follow manufacturer’s instructions and supply students with a photograph of the ladder with molecular weights labeled.
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LAB SESSION 4 Note to prep staff: Pool students’ pEGFP-N1 and perform the NcoI/NotI digests. Scaling-up the reaction in a large tube and then aliquoting from this is much easier than setting up individual reactions. Incubating the digests overnight a day or two before the lab works well. Always run a small amount on a gel to ensure complete digestion. The students will use the following supplies: ●
●
● ● ●
100 µl pEGFP-N1 (30 µg) digested with NcoI and NotI by the instructor as described in Lab Session 4.II.A. (This pEGFP-N1 DNA was collected from the class at the end of Lab Session 2.) 4 µl low DNA mass ladder (Invitrogen, catalog number 10068-013) (or λ HindIII NEB catalog number N3012S: follow manufacturer’s instructions) Razor blade or scalpel 1 mg/ml Crystal Violet (enough for 300 µl per station) Plastic sheets to cut gel on
The following supplies and reagents are included in the Qbiogene BIO 101® Geneclean® II kit (catalog number 1001-400). Aliquot the following amounts for student use, but have extra aliquots handy in case large gel slices are extracted. ● ● ● ●
Glass milk (12 µl aliquoted per group) NEW wash (600 µl aliquoted per group) NaI solution TBE modifying buffer
LAB SESSION 5 Note to prep staff: 1. Two days before the lab period, streak an E. coli expression host strain such as BL21(DE3) onto LB agar (no antibiotics). 2. One night before the lab period, inoculate this E. coli into 2 ml of LB medium (no antibiotics) for each group. Be careful to use aseptic technique because there are no antibiotics in the medium. The students will use the following supplies: ● ● ● ● ● ● ● ● ●
Spectrophotometer cuvettes Transformation and storage solution (TSS) (10 ml per group) SOC broth (2 ml per group) Supercoiled pET-41a(+) DNA at 10 ng/µl (aliquot 2 µl per group) 100 ml LB broth in 1 L flask per group LB agar containing 50 µg/ml kanamycin (LB/kan plates) (2 plates per group) Dry ice 95% ethanol for dry ice bath Microcentrifuge tube holder or cheesecloth for freezing cells
Recipes Transformation and Storage Solution (TSS) Ingredient
Amount
Tryptone Yeast extract NaCl Polyethylene glycol, MW 3350 or 6000–8000
0.10 g 0.05 g 0.10 g 10% w/v
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Ingredient
Amount
MgCl2. 6H20 DMSO Deionized water
0.10 g 5% v/v 9 ml
Bring to pH 6.5, add deionized water to 10 ml and filter sterilize. Store at 4˚C SOC Broth SOC is a richer medium than LB and is frequently used to help E. coli recover from transformation. Prepare SOC in three steps: 1. Prepare 2 M magnesium stock by combining 1 M MgCl2 and 1 M MgSO4; filter sterilize and store at room temperature. 2. Prepare 2 M glucose stock; filter sterilize and store at −20˚C. 3. Assemble SOC medium. For 100 ml: ● ● ● ●
Bacto Tryptone, 2 g Yeast extract, 0.5 g NaCl, 10 mM KCl, 2.5 mM
Bring to 1 liter with deionized water and autoclave. When cool, add 1 ml of magnesium stock and 1 ml of glucose stock.
LAB SESSION 6 The students will use the following supplies: ●
Vector and insert DNA Students should have digested and undigested vector and isolated egfp insert saved in their freezer boxes from previous labs. They should have the calculations they made at the end of Lab Session 4.
●
● ● ● ●
5 µl T4 DNA ligase per group. NOTE: This protocol uses NEB (New England Biolabs) T4 DNA ligase, catalog number M0202L. If using a different brand of T4 DNA ligase, follow manufacturer’s instructions. The incubation time may be significantly longer. 8 µl ligase buffer per group 500 µl LB or SOC per group 6 LB/kan plates per group 150 µl competent E. coli BL21(DE3) cells per group Thaw frozen compentent cells immediately before use. Cells should not sit on ice for more than a few minutes. Cells should not sit at room temperature for any period of time.
● ● ● ●
Sterile water 10× DNA loading buffer 1 kb ladder (Invitrogen) Glass plate spreaders
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LAB SESSION 7A: INTERIM LAB PERIOD Note to prep staff: The afternoon before the interim lab, streak out the following cultures on several LB/kan plates each: 1. pAD1 in host strain BL21(DE3)(positive control) 2. pET-41a(+) in host strain BL21(DE3)(negative control) 3. Mixture of pAD1 and pET-41a(+) in host strain BL21(DE3) (mixed unknowns) Streak in quadrants in order to assure isolated colonies. The students will each need a colony of 1 and 2 for their positive and negative controls, respectively. The mixed unknowns will be used as back-up by students who did not obtain any transformants on their own plates. The students will use the following supplies: ● ●
● ● ●
●
●
3 LB/kanamycin plates per group 3 grid stickers per group (Diversified Biotech PetriSticker™ 50 square grid OR use grids in Figure 7.1) Sterile toothpicks for each group Several LB/kan plates containing pAD1 in host strain BL21(DE3)(positive control) Several LB/kan plates containing pET-41a(+) in host strain BL21(DE3)(negative control) Several LB/kan plates containing a mixture of pAD1 and pET-41a(+) in host strain BL21(DE3) (mixed unknowns) Students’ transformation plates from Lab Session 6
LAB SESSION 7A: REGULAR LAB PERIOD Note to prep staff: 1. Place students’ replica plates inverted in the refrigerator (lids down) until the afternoon prior to the lab session. 2. The afternoon prior to the lab session, place students’ replica plates inverted in the 37˚C incubator. 3. The morning of the lab session, 3–4 hours prior to the beginning of lab, begin the colony lift for the monoclonal antibody probe as described in Lab Session 7C, steps 1–5.
LAB SESSION 7B The students will use the following supplies: ● ● ● ● ● ● ● ● ●
1 circular nylon membrane filter per group 4 pieces Whatman 3MM paper (to sandwich filter) per group 2 ml denaturing solution per group 2 ml neutralizing solution per group 1 ml 2× SSC per group 0.6 ml proteinase K (2 mg/ml) per group Plastic wrap Empty Petri dishes for washes Forceps For the dig-labeling reaction, you will need:
●
100 mM stocks of each: dATP, dCTP, dGTP, dTTP
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1 mM stock of dig-dUTP pEGFP-N1 (for DNA template) Taq DNA polymerase PCR buffer with magnesium (10X stock) 100 pmol/µl stock of primer 1 (CTTGTACAGCTCGTCCATGC) 100 pmol/µl stock of primer 2 (AGAGTCCCATGGTGAGCAAG)
Recipes Denaturing Solution Compound
Final Concentration
Amount Needed for 20 Groups
0.5 M 1.5 M
25 ml 150 ml to 500 ml
NaOH, 10 N NaCl, 5 M Deionized water Neutralizing Solution (0.5 M TRIS pH 7.5, 1.5 M NaCl) Compound Tris (pH 7.5), 1 M NaCl, 5 M Deionized water
Final Concentration
Amount Needed for 20 Groups
0.5 M 1.5 M
250 ml 150 ml to 500 ml
20× SSC Compound NaCl Sodium citrate Deionized water
Final Concentration
Amount Needed for 1 Liter
3.0 M 0.3 M
175.3 g 88.2 g to 1 liter
Adjust pH to 7.0 using 14 N HCl. 2× SSC Dilute 20× SSC 1:10 with water.
LAB SESSION 7C The students will use the following supplies: ● ● ● ● ●
● ● ● ● ● ● ●
Nitrocellulose (1 circle per group) (used by prep staff) IPTG solution (20 mg/ml in H20) (20 ml, used by prep staff) LB/kan plates (1 per group) (used by prep staff) Chloroform Glass Petri plates with support mesh (or toothpicks taped to the edges to hold plastic Petri dishes above) Lysis buffer (8 ml/group) Blocking solution (10ml/group) 1× wash buffer/IGEPAL/milk (10 ml/group) 1× wash buffer/ milk (10 ml/group) 3 squares Whatman paper/group Empty Petri dishes Pencil
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Recipes Blocking Solution Combine the following and adjust to pH 7.5. Aliquot 10 ml into 15-ml conical tubes, one per group, and freeze at −20˚C. Note: Extra blocking solution will be needed to make Lysis buffer.
Component *
Nonfat powdered milk NaCl, 1 M Tris (pH 8.0), 1 M Deionized water
Amount Added
Final Concentration
15 g 18.75 ml 6.25 ml to 500 ml
3% 75 mM 25 mM
*
Nonfat powdered milk is available at most grocery stores. Bovine serum albumin (BSA; fraction V) can be substituted for nonfat powdered milk.
Lysis Buffer Aliquot 8 ml into 15-ml conical tubes, one per group. Store at −20˚C. Blocking solution Lysozyme DNase (1 mg/ml )* MgCl2, 1 M
250 mls 10 mg 25 fl 1.25 ml
Pancreatic DNase (1 mg/ml), prepare in deionized, sterile water and store at −20˚C. Use to make lysis buffer. *
1× Wash Buffer/Milk and 1× Wash Buffer/IGEPAL*/Milk *
IGEPAL is a nonionic, non-denaturing detergent that is also known by the following names: Nonidet P40; NP 40; IGEPAL CA-630; and Nonylphenyl-polyethylene glycol. For both recipes, first make a 10× stock: 10× Wash Buffer Tris (pH 7.4), 1 M NaCl Deionized water
200 ml 87.6 g to 1 liter
1× Wash Buffer/Milk Prepare 100 ml aliquots for general use by the students or aliquot 10 ml per group. Component 10× wash buffer Deionized water Nonfat powdered milk
Amount Added
Final Concentration
100 ml 900 ml 1g
1× 0.1%
1× Wash Buffer/IGEPAL/Milk Use a 1 ml syringe to add 0.5 ml IGEPAL to 1 L of “1× wash buffer/ milk” (this is a 0.05% final concentration of IGEPAL). Prepare 100 ml aliquots for general use by the students or aliquot 10 ml per group.
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LAB SESSION 8 The students will use the following supplies: ●
● ● ● ● ● ● ● ●
● ● ● ● ●
Hybridization buffer (250 ml total) (We use DIG Easy Hyb from Roche: catalog number 1 603 558.) DIG-labeled egfp probe (prepared in Lab Session 7) 0.5× wash solution (enough for 60 ml/group, pre-warmed to 65˚ C) 2× wash solution (enough for 60 ml/group, room temperature) l× wash buffer/milk (60 ml/group) (same recipe as in Lab Session 7C) α dig-HRP antibody (4 µl/group) (Pierce Biotechnology product number 31468) Blocking solution (30 ml/group) (same recipe as in Lab Session 7C) Peroxide stain (from chloronaphthol stock, 7.5 ml/group) Washing buffer (catalog number 1 585 762 from Roche as part of the DIG Wash and Block Buffer Set). This comes as a 10× solution. Dilute 10-fold before distributing. Empty Petri dishes Paper towels LB/kan plate (1 per group) Grid sticker (1 per group) Sterile toothpicks
Recipes 20× SSC Prepare as described in Lab Session 7B. SDS (10%) Electrophoresis-grade sodium dodecyl (also called lauryl) sulfate 50 g Deionized water to 500 ml Wear a mask when weighing SDS. Do not autoclave SDS. 2× Wash Solution (2× SSC Containing 0.1% SDS) 20× SSC 10% SDS Deionized water
200 ml 20 ml to 2 liters
0.5× Wash Solution (0.5× SSC Containing 0.1% SDS) 20× SSC 10% SDS Deionized water
50 ml 20 ml to 2 liters
Chloronaphthol Stock Solution This is used by the staff to make peroxide stain. Note: Wear gloves—chloronaphthol is a suspected carcinogen. Combine the following and store at −20˚C in a light-tight bottle: 4-Chloro-1-naphthol Ice-cold methanol
150 mg (0.3% final concentration) 50 ml
Peroxide Stain This should be made fresh directly before lab. Purchase new hydrogen peroxide each semester.
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Component
Amount Added
Final Concentration
17 ml 68 ml 5 ml 10 ml 35 µ1
0.05%
Chloronaphthol stock Deionized water Tris (pH 7.4), 1 M NaCl, 1 M H2O2,*30%
50 mM 100 mM 0.02%
*
Hydrogen peroxide (H2O2) comes as a 30% solution and is stored at 4˚C
LAB SESSION 9A The students will use the following supplies: ● 1× wash buffer/IGEPAL/milk (15 ml/group) (same recipe as in Lab Session 7C) ● 1× wash buffer/ milk (70 ml/group) (same recipe as in Lab Session 7C) ● α-GFP antibody (7.5 µl/group) (Clontech catalog number NC9777966) ● Blocking solution (15 ml/group) (same recipe as in Lab Session 7C) ● Goat anti-mouse conjugated to peroxidase (GAMP) (15 µl/group) (Sigma-Aldrich catalog number A-3673) ● Peroxide stain (from chloronaphthol stock, 7.5 ml/group) (same recipe as in Lab Session 8) ● Empty Petri dishes ● Paper towels
LAB SESSION 9B The students will use the following supplies: ● ●
1 set of PCR strip tubes and caps per group PCR master mix (180 µl per group)
Item Water A (100 mM stock) T (100 mM stock) C (100 mM stock) G (100 mM stock) Primer 1 100 pmol/µl (pad1sense)* Primer 2 100 pmol/µl (pad1anti)* 10× Buffer with Mg Taq polymerase
per 180 µl 155.2 µl 0.36 µl 0.36 µl 0.36 µl 0.36 µl 0.9 µl 0.9 µl 18 µl 3.6 µl
*
Note that Primer 2 used for this experiment differs from the Primer 2 that was used in the dig-labeling reaction of lab session 7b. Be sure to use the correct primer.
● ●
Primer1: CTTGTACAGCTCGTCCATGC. Primer2: CAAGCTACCTGAAATGCTGA.
LAB SESSION 9C The students will use the following supplies: ●
LB/kanamycin/IPTG plates (1 per group): 1 ml of 20 mg/ml IPTG and 1 ml of 50 mg/ml kanamycin is added to 1 liter of cooled LB agar. Swirl then pour plates.
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●
LB/kanamycin (1 per group): 1 ml of 50 mg/ml kanamycin is added to 1 liter of cooled LB agar. Swirl then pour plates. Sterile toothpicks
LAB SESSION 10A: INTERIM LAB The students will use the following supplies: ● ● ●
Six 2 ml aliquots of LB/kan broth per group (in snap-cap culture tubes) Sterile toothpicks Access to their replica plate (replicated master plate on LB/kan from Lab Session 9C)
LAB SESSION 10C The students will use the following supplies: ● ● ● ● ● ● ● ● ● ● ●
● ●
Miniprep solution I (700 µl/group) Miniprep solution II (1.4 ml/group) Miniprep solution III (1 ml/group) 95% ETOH, stored at −20˚C 70% ETOH, room temperature RNase (20 mg/ml) (10 µl/group) Not I (4 µl/group) Nco I (8 µl/group) BSA (4 µl/group) (supplied as a 10 mg/ml stock, which is 100X) 1× TE buffer 10× restriction buffer 3 (36 µl/group): This is for NEB enzymes and buffers. Follow manufacturer’s instructions if you are using a different brand. Sterile deionized water 10× DNA loading buffer (students should have some saved from previous labs)
Recipes Miniprep Solution I Component Glucose Tris-HCl (pH 8.0), 1 M EDTA, 0.5 M
Final Concentration
Amount Needed for 100 ml
50 mM 25 mM 10 mM
1.81 g 5 ml 0.4 ml
Miniprep Solution II Make fresh with stocks of NaOH and sodium dodecyl sulfate (SDS): Component NaOH, 10 N SDS, 10% Deionized water
Final Concentration
Amount Needed for 200 ml
0.2 N 1%
4 ml 20 ml 176 ml
Miniprep Solution III For 100 ml: Potassium acetate, 5 M Glacial acetic acid Deionized water
60 ml 11.5 ml 28.5 ml
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RNase (20 mg/ml) For 1 ml combine the following: Tris (pH 8.0), 1 M NaCl, 5 M RNase (pancreatic or RNase A)
10 µl 3 µl 20 mg
Boil for 15 min. This step is critical for inactivating any contaminating DNases present Store at −20˚C. Aliquot 10 µl into microcentrifuge tubes, one per group.
LAB SESSION 11A: INTERIM LAB The students will use the following supplies: ● ● ● ● ●
Snap-cap tubes with 1 ml LB/kan (3 per group) Empty snap-cap tubes (3 per group) Sterile toothpicks Sharpie markers Students’ master plates (the most freshly streaked ones)
LAB SESSION 11B: REGULAR LAB PERIOD Note to prep staff: 1. Two days before the laboratory: streak LB/kan plates of the positive and negative control strains. ●
positive: pAD1 in E. coli strain BL21(DE3) or other appropriate expression strain
●
negative: pET-41a(+) in E. coli strain BL21(DE3) or other appropriate expression strain
2. One day before lab: incubate the students’ LB/kan broth cultures overnight at 37˚C. Also inoculate and incubate positive and negative controls into LB/kan broth. The positive and negative controls are the pAD1 and pET-41a(+) plasmids respectively, in E. coli host strain BL21(DE3). 3. The morning of the lab: 3–4 hours before the start of lab, subculture 1 ml of each student culture into 2 ml 2× YT medium containing kanamycin and IPTG, as described in Lab Session 11B, Section IIA, steps 1–3. Also subculture the positive and negative control into 2× YT/kan/IPTG. 4. Fill and freeze ice blocks for electroblotting prior to each day’s lab. Prep staff use: ●
2× YT broth with kanamycin and IPTG The students will use the following supplies:
●
●
● ● ●
pAD1 and pET-41a(+) E. coli BL21(DE3) cultures that have been incubated in 2× YT/kan/IPTG 3–4 hours before lab Student cultures that have been incubated in 2× YT/kan/IPTG 3–4 hours before lab Locking or screw-cap microfuge tubes (5 per group) 1× Tris-glycine running buffer (6 liters) IPTG (20 mg/ml stock to add to medium)
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● ●
● ● ● ● ● ● ● ●
● ● ● ● ●
Polyacrylamide gel (1 per group) (Use a gel with a polyacrylamide concentration in the range of 7.5% to 10%, such as Bio-Rad catalog numbers 161-1210 or 1611100, or ISC BioExpress catalog numbers E-4325-010 or E-4326-008.) 2× sample (loading) buffer (1 ml/group) MW marker, 12 µl per group (NEB broad range molecular weight marker, catalog number P7702S) Disposable square Petri dishes (Fisher 08-757-11A) Whatman (filter) paper (2 pieces per group, cut to the size of the fiber pads) Nitrocellulose (1 piece per group, cut to the size of the gels) Fiber pads (2 per group) (NOT disposable) TBS-T (40 ml per group) Blocking solution (TBS-T plus 5% powdered milk) (10 ml per group) Transfer buffer, 5× stock (estimate 200 ml per lab station) Ponceau S stain (Sigma-Aldrich catalog number P7170), approximately 10 ml per group Ponceau S stain may be saved and reused. Razor blades Methanol (estimate 200 ml per lab station) Glass casserole dishes Glass tubes or Pasteur pipets to use as “rolling pins” Cap locks to prevent microcentrifuge tube lids from popping open during heating (Fisher catalog number NC9346739)
Recipes IPTG (20 mg/ml) Dissolve 0.l g IPTG in 5 ml deionized water. Filter-sterilize. This is a 1000× stock. Store at −20˚C. 2× YT Broth Component Amount Needed for 20 Groups Bacto tryptone Yeast extract NaCl Deionized water
16 g 10 g 5g to 1 liter
Autoclave and store at room temperature. 2× YT/Kan/IPTG To 100 ml 2× YT, add 100 µl kanamycin (50 mg/ml stock) and 100 µl IPTG (20 mg/ml stock). The final concentration of kanamycin is 50 µg/ml, and the final concentration of IPTG is 20 µg/ml. Tris–Glycine Running Buffer Recipes for the 1× concentration and a 10× stock solution are below: Component
Tris base Glycine SDS Deionized H2O
Amount
Final Concentration
1×
10×
1×
10×
6.0 g 28.8 g 2.0 g to 2000 ml
30 g 141 g 10 g to 1000 ml
25 mM 190 mM 0.1%
0.25 M 1.9 M 1%
Place carboys of 1× running buffer in the lab. It is a good idea to keep extra 10× stock on hand.
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2× Sample (Loading) Buffer Prepare under the fume hood: Component
Amount
Final Concentration
SDS Tris (pH 6.8), 0.5 M Sucrose β-Mercaptoethanol EDTA (pH 7.0), 0.1 M Bromphenol blue, 1% Deionized H2Oto
1g 25 ml 15 g 10 ml 1 ml 5 ml 100 ml
1% 125 mM 15% 10% 1 mM 0.05%
Aliquot into 1-ml volumes (one per group) and store frozen TBS (Tris-Buffered Saline, pH 7.6) Component
Amount
Final Concentration
Tris base NaCl HCl (1 M), to pH 7.6 Deionized water
2.42 g 8g 3.8 ml to 1 liter
20 mM 137 mM
TBS-T 0.1% Tween 20 in TBS. Prepare 40-ml aliquots or prepare a large amount for common use. Blocking Solution TBS-T plus 5% nonfat milk powder (or bovine serum albumin fraction V). Store at −20˚C. 5× Transfer Buffer To prepare 1 liter, add: Component
Amount
Final Concentration
Tris base Glycine SDS Deionized water
15.15 g 72.05 g 5g to 1 liter
0.125 M 0.95 M 0.5%
Store 5× transfer buffer in carboys at room temperature. Students will prepare 1× transfer buffer directly before use, by mixing 200 ml 5× transfer buffer, 200 ml methanol, and 600 ml deionized water. A large bottle of methanol should be available in the fume hood for student use. After use, the 1× buffer containing methanol should be poured in a methanol waste container and discarded according to the safety regulations at your institution.
LAB SESSION 12 The students will use the following supplies: ● ● ●
TBS-T (same recipe as in Lab Session 11) α-GFP antibody (8.5 µl/group) (Clontech catalog number NC9777966) Goat α-mouse antibody conjugated to peroxidase (GAMP) (16 µl/group) (SigmaAldrich catalog number A-3673)
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● ● ●
Peroxide stain (made fresh from chloronaphthol stock, 10 ml/group) (same recipe as in Session 8) 30% hydrogen peroxide should be less than 1 month old LB/kan plate (1 per group) Sterile toothpicks Whatman (filter) paper (2 pieces per group, cut to the size of the fiber pads)
LAB SESSION 13A: INTERIM LAB The students will use the following supplies: ● ● ● ●
Sterile toothpicks LB/kan 2 ml aliquots in snap-cap culture tubes (1 per group) Sharpie markers Students’ replica plates of positive clone (streaked last lab session)
LAB SESSION 13B: REGULAR LAB PERIOD Note to prep staff: 1. The evening before lab: incubate students’ cultures (that were stored in the refrigerator after the interim lab) overnight shaking at 37˚C. Also inoculate and incubate E. coli strain BL21(DE3) harboring the pAD1 plasmid. This can be used as a backup for purification if students accidentally spill their own cultures. 2. The morning of lab: subculture students’ 2 ml cultures and backup culture into 250 ml flasks containing 100 ml 2× YT/Kan/IPTG each. Incubate shaking at 37˚ C for 3–4 hours prior to the start of lab. The students will use the following supplies: ● ●
●
● ● ● ●
● ●
●
● ●
Oak Ridge tubes (2 per group) 2× YT/kan/IPTG, 100 ml/group (in 250 ml flasks for prep staff to use for subculturing) 2× YT or LB for resuspending bacteria (a few small bottles per lab or one 1.3 ml tube per group) 2× sample buffer (if needed: students likely have enough left from last week) Lysozyme (10 mg/ml), 20 µl per group Dry ice GST-Bind resin (2 ml settled bead volume in 15 ml conical tube, 1 per group) (Novagen catalog number 70541-4, or equivalent GST affinity resin) Chromatography column/stopcock (1 per group) Pefabloc® (4-(2-Aminoethyl)benzenesulfonyl fluoride hydrochloride, AEBSF, Sigma-Aldrich catalog number 76307), 0.1 M stock solution (prepare stock solution with sterile, deionized water, and freeze at −20˚C) GST Bind/Wash buffer, room temp (34 ml per group) (a component of the Novagen GST-Bind Buffer kit, catalog number 70534-3, or equivalent): Dilute 10× stock according to manufacturer’s instructions. GST Bind/Wash buffer with Pefabloc, ice-cold (0.75 ml per group) GST elution buffer (7 ml per group) (a component of the Novagen GST-Bind Buffer kit, catalog number 70534-3, or equivalent): Prepare from 10× Glutathione Reconstitution Buffer and reduced glutathione, according to manufacturer’s instructions.
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Recipes GST Bind/Wash Buffer with Pefabloc®* Add 1 ml of the 0.1 M Pefabloc® stock solution to 50 ml 1× Bind/Wash buffer. Mix and keep at 4˚ C until used.
LAB SESSION 14 The students will use the following supplies: ● ● ●
● ● ● ● ●
● ● ● ●
*
Sterile H2O BSA (1 mg/ml) (100 µl per group) Coomassie G250/Bradford Reagent (diluted if specified by the manufacturer), 7.5 ml per group (Sigma-Aldrich catalog number B6916) 96 well plate (1 per group) Multichannel pipetters and multichanel resevoirs, if available Polyacrylamide gel (1 per group, as described in Lab Session 11) 2× sample (loading) buffer Broad range molecular weight marker for SDS-PAGE (Bio-Rad catalog number 161-0317) 1X Tris-glycine running buffer (6 liters, recipe as described in Lab Session 11) Gelcode Blue staining reagent, 20 ml per group (Pierce catalog number 24590) Square Petri dish (1 per group) Cap locks to prevent microcentrifuge tube lids from popping open during heating (Fisher catalog number NC9346739)
Pefabloc® is a protease inhibitor. Wear gloves, eye protection, and lab coat when working with this chemical.
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APPENDIX
3 Making Sense of Orientation hen making a fusion protein, it is important to maintain the correct reading frame. In the following example, an SphI fragment corresponding to a calcium-binding peptide (cbp) (Figure A3.1) is ligated to a site at the 3′ end of the gfp gene. The fragment can be ligated in two orientations because it has the SphI cohesive ends on both sides. In one orientation, the reading frame is maintained while in the other orientation, a stop codon is found just after the SphI site.
W
SphI 5’ GCATGCCGATCTAGAGCTCGTCGTGCTTCTCATCATCAGAATCNTTGCCATCCTCGGCATCAGAGTCGGCCTTGTCCTCGTCTGCCTTCT 3’ CGTACGGCTAGATCTCGAGCAGCACGAAGAGTAGTAGTCTTAGNAACGGTAGGAGCCGTAGTCTCAGCCGGAACAGGAGCAGACGGAAGA CATCGTCTTCCTCATCCTCTAGGTCATCATCCTCATCATCCCCACCCTTGGCGGCATCCTCTTCTTCCTTCTTTTTCTCGGCCTCATCAA GTAGCAGAAGGAGTAGGAGATCCAGTAGTAGGAGTAGTAGGGGTGGGAACCGCCGTAGGAGAAGAAGGAAGAAAAAGAGCCGGAGTAGTT AAGCAGCCTTTTCTGCCTCCTTGTGCTTGCCCCAGGTCTCCTCTGCAAAAGTCTTGGCCAACGCAGGGTCATCAGTGATGATGATGTTGT TTCGTCGGAAAAGACGGAGGAACACGAACGGGGTCCAGAGGAGACGTTTTCAGAACCGGTTGCGTCCCAGTAGTCACTACTACTACAACA CGAACAGAGTGCCCGATTTAACCTGCCACAGCTCAATGCCAATGTACTTCAAGCTGTCGAAGGCGTAAATGTATGGATCATCCTTAAAAT GCTTGTCTCACGGGCTAAATTGGACGGTGTCGAGTTACGGTTACATGAAGTTCGACAGCTTCCGCATTTACATACCTAGTAGGAATTTTA SphI CTGGGTTGTCAATCATACGGGCATGC 3’ GACCCAACAGTTAGTATGCCCGTACG 5’
Fig. A3.1 Nucleotide sequence of the SphI fragment containing the cbp gene. The sequence is shown in the reverse orientation, with the upper strand corresponding to the antisense strand, and the lower strand corresponding to the sense strand. Note that if the DNA were transcribed in this orientation, a stop codon, TAG, could be located just downstream of the SphI site.
In the correct orientation, the top strand of the SphI fragment is in frame with gfp. In the incorrect orientation, the bottom strand, beginning at the 5′ end, is adjacent to the gfp sequence. In the two examples below, the amino acid sequence is shown directly beneath the DNA sequence. The SphI sites are bold and underlined (6-bp palindromic sites). The stop codon is shown in bold (TAG). First example: cbp fragment cloned in the correct orientation: Second example: cbp fragment cloned in the incorrect orientation:
Thought Question What is the probability of encountering a stop codon by chance? Use the probability formula for independent events, 1/4n and remember there are three stop codons.
143
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promoter
gfp
gfp::cbp correct orientation
SphI cbp SphI terminator
Fig. A3.2 Plasmid DNA containing a plant promoter, gfp open reading frame fused to the calcium binding peptide (cbp) open reading frame, and plant terminator sequence. Bacterial genes are not shown for simplicity.
ATGACGCACAATCCCACTATCCTTCGCAAGACCCTTCCTCTATATAAGGAAGTTCATTTCATTTGGAGATCTAGAGGATCCTCTAGAATG MetThrHisAsnProThrIleLeuArgLysThrLeuProLeuTyrLysGluValHisPheIleTrpArgSerArgGlySerSerArgMet AAGACTAATCTTTTTCTCTTTCTCATCTTTTCACTTCTCCTATCATTATCCTCGGCCGAATTCAGTAAAGGAGAAGAACTTTTCACTGGA LysThrAsnLeuPheLeuPheLeuIlePheSerLeuLeuLeuSerLeuSerSerAlaGluPheSerLysGlyGluGluLeuPheThrGly GTTGTCCCAATTCTTGTTGAATTAGATGGTGATGTTAATGGGCACAAATTTTCTGTCAGTGGAGAGGGTGAAGGTGATGCAACATACGGA ValValProIleLeuValGluLeuAspGlyAspValAsnGlyHisLysPheSerValSerGlyGluGlyGluGlyAspAlaThrTyrGly AAACTTACCCTTAAATTTATTTGCACTACTGGAAAACTACCTGTTCCATGGCCAACACTTGTCACTACTTTCTCTTATGGTGTTCAATGC LysLeuThrLeuLysPheIleCysThrThrGlyLysLeuProValProTrpProThrLeuValThrThrPheSerTyrGlyValGlnCys TTTTCAAGATACCCAGATCATATGAAGCGGCACGACTTCTTCAAGAGCGCCATGCCTGAGGGATACGTGCAGGAGAGGACCATCTTCTTC PheSerArgTyrProAspHisMetLysArgHisAspPhePheLysSerAlaMetProGluGlyTyrValGlnGluArgThrIlePhePhe AAGGACGACGGGAACTACAAGACACGTGCTGAAGTCAAGTTTGAGGGAGACACCCTCGTCAACAGGATCGAGCTTAAGGGAATCGATTTC LysAspAspGlyAsnTyrLysThrArgAlaGluValLysPheGluGlyAspThrLeuValAsnArgIleGluLeuLysGlyIleAspPhe AAGGAGGACGGAAACATCCTCGGCCACAAGTTGGAATACAACTACAACTCCCACAACGTATACATCATGGCCGACAAGCAAAAGAACGGC LysGluAspGlyAsnIleLeuGlyHisLysLeuGluTyrAsnTyrAsnSerHisAsnValTyrIleMetAlaAspLysGlnLysAsnGly ATCAAAGCCAACTTCAAGACCCGCCACAACATCGAAGACGGCGGCGTGCAACTCGCTGATCATTATCAACAAAATACTCCAATTGGCGAT IleLysAlaAsnPheLysThrArgHisAsnIleGluAspGlyGlyValGlnLeuAlaAspHisTyrGlnGlnAsnThrProIleGlyAsp GGCCCTGTCCTTTTACCAGACAACCATTACCTGTCCACACAATCTGCCCTTTCGAAAGATCCCAACGAAAAGAGAGACCACATGGTCCTT GlyProValLeuLeuProAspAsnHisTyrLeuSerThrGlnSerAlaLeuSerLysAspProAsnGluLysArgAspHisMetValLeu SphI CTTGAGTTTGTAACAGCTGCTGGGATTACACATGGCATGGATGAACTATACAAACAGCATGCCCGTATGATTGACAACCCAGATTTTAAG LeuGluPheValThrAlaAlaGlyIleThrHisGlyMetAspGluLeuTyrLysGlnHisAlaArgMetIleAspAsnProAspPheLys GATGATCCATACATTTACGCCTTCGACAGCTTGAAGTACATTGGCATTGAGCTGTGGCAGGTTAAATCGGGCACTCTGTTCGACAACATC AspAspProTyrIleTyrAlaPheAspSerLeuLysTyrIleGlyIleGluLeuTrpGlnValLysSerGlyThrLeuPheAspAsnIle ATCATCACTGATGACCCTGCGTTGGCCAAGACTTTTGCAGAGGAGACCTGGGGCAAGCACAAGGAGGCAGAAAAGGCTGCTTTTGATGAG IleIleThrAspAspProAlaLeuAlaLysThrPheAlaGluGluThrTrpGlyLysHisLysGluAlaGluLysAlaAlaPheAspGlu GCCGAGAAAAAGAAGGAAGAAGAGGATGCCGCCAAGGGTGGGGATGATGAGGATGATGACCTAGAGGATGAGGAAGACGATGAGAAGGCA AlaGluLysLysLysGluGluGluAspAlaAlaLysGlyGlyAspAspGluAspAspAspLeuGluAspGluGluAspAspGluLysAla SphI GACGAGGACAAGGCCGACTCTGATGCCGAGGATGGCAGAGATTCTGATGATGAGAAGCACGACGAGCTCTAGATCGGCATGCCCTGCTTT AspGluAspLysAlaAspSerAspAlaGluAspGlyArgAspSerAspAspGluLysHisAspGluLeu AATGAGATATGCGAGACGCCTATGATCGCATGATATTTGCTTTCAATTCTGTTGTGCACGTTGTAA
Fig. A3.3
Nucleotide and amino acid sequence of the gfp::cbp fusion
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Appendix 3: Making Sense of Orientation
promoter
gfp
gfp::cbp incorrect orientation
SphI cbp SphI terminator
Fig. A3.4 Plasmid DNA with the same cbp open reading frame as in A3.2, but in the opposite orientation. In this case only the gfp open reading frame is translated.
ATGACGCACAATCCCACTATCCTTCGCAAGACCCTTCCTCTATATAAGGAAGTTCATTTCATTTGGAGATCTAGAGGATCCTCTAGAATG MetThrHisAsnProThrIleLeuArgLysThrLeuProLeuTyrLysGluValHisPheIleTrpArgSerArgGlySerSerArgMet AAGACTAATCTTTTTCTCTTTCTCATCTTTTCACTTCTCCTATCATTATCCTCGGCCGAATTCAGTAAAGGAGAAGAACTTTTCACTGGA LysThrAsnLeuPheLeuPheLeuIlePheSerLeuLeuLeuSerLeuSerSerAlaGluPheSerLysGlyGluGluLeuPheThrGly GTTGTCCCAATTCTTGTTGAATTAGATGGTGATGTTAATGGGCACAAATTTTCTGTCAGTGGAGAGGGTGAAGGTGATGCAACATACGGA ValValProIleLeuValGluLeuAspGlyAspValAsnGlyHisLysPheSerValSerGlyGluGlyGluGlyAspAlaThrTyrGly AAACTTACCCTTAAATTTATTTGCACTACTGGAAAACTACCTGTTCCATGGCCAACACTTGTCACTACTTTCTCTTATGGTGTTCAATGC LysLeuThrLeuLysPheIleCysThrThrGlyLysLeuProValProTrpProThrLeuValThrThrPheSerTyrGlyValGlnCys TTTTCAAGATACCCAGATCATATGAAGCGGCACGACTTCTTCAAGAGCGCCATGCCTGAGGGATACGTGCAGGAGAGGACCATCTTCTTC PheSerArgTyrProAspHisMetLysArgHisAspPhePheLysSerAlaMetProGluGlyTyrValGlnGluArgThrIlePhePhe AAGGACGACGGGAACTACAAGACACGTGCTGAAGTCAAGTTTGAGGGAGACACCCTCGTCAACAGGATCGAGCTTAAGGGAATCGATTTC LysAspAspGlyAsnTyrLysThrArgAlaGluValLysPheGluGlyAspThrLeuValAsnArgIleGluLeuLysGlyIleAspPhe AAGGAGGACGGAAACATCCTCGGCCACAAGTTGGAATACAACTACAACTCCCACAACGTATACATCATGGCCGACAAGCAAAAGAACGGC LysGluAspGlyAsnIleLeuGlyHisLysLeuGluTyrAsnTyrAsnSerHisAsnValTyrIleMetAlaAspLysGlnLysAsnGly ATCAAAGCCAACTTCAAGACCCGCCACAACATCGAAGACGGCGGCGTGCAACTCGCTGATCATTATCAACAAAATACTCCAATTGGCGAT IleLysAlaAsnPheLysThrArgHisAsnIleGluAspGlyGlyValGlnLeuAlaAspHisTyrGlnGlnAsnThrProIleGlyAsp GGCCCTGTCCTTTTACCAGACAACCATTACCTGTCCACACAATCTGCCCTTTCGAAAGATCCCAACGAAAAGAGAGACCACATGGTCCTT GlyProValLeuLeuProAspAsnHisTyrLeuSerThrGlnSerAlaLeuSerLysAspProAsnGluLysArgAspHisMetValLeu SphI CTTGAGTTTGTAACAGCTGCTGGGATTACACATGGCATGGATGAACTATACAAACAGCATGGCATGCCGATCTAGGTCGTGCTTCTCATC LeuGluPheValThrAlaAlaGlyIleThrHisGlyMetAspGluLeuTyrLysGlnHisGlyMetProIle ATCAGAATCNTTGCCATCCTCGGCATCAGAGTCGGCCTTGTCCTCGTCTGCCTTCTCATCGTCTTCCTCATCCTCTAGGTCATCATCCTC ATCATCCCCACCCTTGGCGGCATCCTCTTCTTCCTTCTTTTTCTCGGCCTCATCAAAAGCAGCCTTTTCTGCCTCCTTGTGCTTGCCCCA GGTCTCCTCTGCAAAAGTCTTGGCCAACGCAGGGTCATCAGTGATGATGATGTTGTCGAACAGAGTGCCCGATTTAACCTGCCACAGCTC SphI AATGCCAATGTACTTCAAGCTGTCGAAGGCGTAAATGTATGGATCATCCTTAAAATCTGGGTTGTCAATCATACGGGCATGCCGCATGCC CGTATGATTGACAACCCAGATTTTAAGGATGATCCATACATTTACGCCTTCGACAGCTTGAAGTACATTGGCATTGA
Fig. A3.5 tein
Nucleotide and amino acid sequence of the defective gfp::cbp fusion pro-
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Index Aequoria victoria, 5, 29 Affinity chromatography, 109 principle steps of, 110 protein purification by, 113–114 Affinity resin biospecific ligand of, 111 composition of, 111f Agarose beads, 111 of dig-labeled egfp probe, 56f egfp DNA isolation from, 30–34 gel, 29, 40 electrophoresis, 24–26 Aliquots, prep list for, 124 Alkaline lysis, 13, 81 for DNA plasmid purification, 14–16 and ethanol precipitation of miniprep DNA, 82–83 Alkaline phosphatase (AP), 54 Amido black, 12 recipe for, 127 Anion exchange chromatography, 13, 26 for plasmid DNA purification, 14–16 Antibiotics, 77 prep list for, 123 Antibodies, 45, 59–60, 95, 115 AP. See Alkaline phosphatase Autoclaving, prep list for, 124–125 Bacterial cells, 13, 109 breaking open, 112–113 lysing of, in colony hybridization with an α-GFP monoclonal antibody, 61 Bacterial suspension cultures for fusion protein purification, 112 growing of, 114 Bacterial waste, prep list for, 124 BamHI, 22
Bicinchoninic acid (BCA) assay, advantages and disadvantages of, 116–117 Biochemically inert spacers, 111 Biospecific ligands, 110–111 Blocking solution, recipe for, 134, 140 Blue-white screening, 81 Bovine serum albumin (BSA), 60, 115 serial dilutions and nitrocellulose spot test, 11–12 Bradford assay, 115 advantages and disadvantages of, 116 composition of assay wells for, 118t of purification fractions, 118–120 Breaking open bacterial cells, 112–113 BSA serial dilutions and nitrocellulose spot test, 11–12 Buffer P3, 15 Buffer QF, 15 Calcium binding peptide (CBP), open reading frames, 144f, 145f Calcium chloride treatment, preparation of transformation-competent cells by, 35–36 CBP. See Calcium binding peptide Chloronaphthol, 54, 60, 101 recipe for, 135 Chromophore regions, 5 Circular DNA, transformation of, 40, 43 Cleaning DNA using spin column, 26–27 Cloning vectors, 19 Colony hybridization, 47, 81 counting transformants, 49 with α-GFP monoclonal antibody probe, 60–62, 67–68 completion, 67–68 flowchart of, 61f
147
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Index Colony hybridization (continued) with DNA probes, 53–56 completion, 63–64 flowchart of, 56f with egfp DNA probe, 54, 57 detection, 64 hybridization, 63–64 prehybridization, 63 stringency washes, 64 with monoclonal antibody probes, 59–62 replica plating, 50–51 Competent cells. See Transformation Coomassie Blue, 96 G-250, 116, 119 Counting transformants, 49 Crude homogenates (CH), removing insoluble debris from, 115 Crystal Violet dye, 30 Denatured DNA, 13, 54 Denaturing solution, recipe for, 133 Density, of water, 11 Destaining solution, recipe for, 127 Digoxigenin-based labeling, 47 of DNA probes, 53–54, 57–58 of egfp, 56f Directional cloning. See Forced cloning Divalent cation-mediated transformation, 42–43 DNA cleaning, using spin column, 26–27 DNA fragments gel separation of, 30–32 ligation of, 39–40 DNA hybridization and detection, principle of, 55f DNA loading buffer, recipe for, 129 DNA probes colony hybridization, 53–56 completion, 63–64 with egfp, 53, 57 flowchart of, 56f digoxigenin labeling of, 53–54 labeling of, by PCR using digoxigenin, 57–58 process of, 53 DNA quantification, 14, 17 EcoRI, 21f egfp DNA isolation from agarose, 30–34 EGFP. See Enhanced green fluorescent protein Electrophoresis. See also Agarose, gel, electrophoresis; Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) of ligation and ligation controls, 44f of ligation reactions, 43–44 PCR sample analysis and gel, 79
of restricted miniprep DNA, 81 on SDS-PAGE and Western Blot, 96–100 types of, 95–96 Emergency contacts, 4 Enhanced green fluorescent protein (EGFP), 5, 26, 47 analytical gel to observe purified, fragment, 33–34 clones expressing, on IPTG, 85–86 DNA probes of, 53–54, 63–64 fluorescence of E. coli expressing, 86f gene, 13, 21–22 isolating, DNA from Agarose, 30–34 isolating, from pEGFP-N1, 29–30 pET-41a preparation and, 19–20 Enzymes, pipetting, 10 Escherichia coli culture, 35–36 fluorescence of, expressing egfp, 86f for plasmid DNA isolation, 13 prep list for, 123 recombinant protein extraction from, 107–116 transformation of, and DNA ligation, 39–41 use of, 5 EtBr. See Ethidium bromide Ethanol precipitation, of miniprep DNA, 82–83 Ethidium bromide (EtBr), 24 recipe for, 127 Experimental procedures, 1 diagram of, 2f Expression vectors, 19, 109 ligation of, 40 orientation, 20–22 reading frame, 22 Fluorescence, 5, 29, 45, 73 Forced cloning, 20f, 21f, 29, 70, 71f Fusion protein, 20–22, 59–61, 89, 91, 95, 101–103, 109–110, 112–113, 115–117 β-Galactosidase, 19–20 GAMP (Goat Anti-Mouse antibody conjugated to Peroxide), 59, 101 Gel sandwich, close-up of, 100f electrophoresis and PCR analysis, 79 separation of DNA fragments, 30–31 GelCode Blue, 96, 117 GENECLEAN® Protocol, 31–32 GeneQuant, 17, 33 gfp. See Green fluorescent protein gfp::cbp fusion protein, 145f nucleotide and amino acid sequence of, 145f Glutathione affinity column, 109–113
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Index Glutathione-S-transferase (gst), 20–22, 109–111 bind/wash buffer recipe, 142 expression of, 20 Green fluorescence assay and preparation of fresh master plate, 73 Green fluorescent protein (gfp), 5, 29 visualization of, 73, 85–86 gst. See Glutathione-S-transferase GST::EGFP fusion protein, 19, 45, 103, 109–111, 115 molecular weight of, 96 purification of, 107 Harvesting IPTG-induced cultures, 112 HindIII, 21f, 39 Horseradish peroxidase (HRP), 54 HRP. See Horseradish peroxidase Hybridization, colony, 47, 49, 81, 95–96, 101 DNA probe, 53–58, 63–64 monoclonal antibody probe, 59–62 with mAB probe, 67–68 Immunoblot. See Western blot Immunological analysis. See Western blot Inoculate cultures for minipreps, 77 for protein purification, 107, 109 for SDS-PAGE, 93 Invitrogen 1 kb ladder, 25f recipe for, 129 Invitrogen low DNA mass ladder, 33f IPTG. See Isopropyl-β-D-thiogalactopyranoside Isopropyl-β-D-thiogalactopyranoside (IPTG), 20, 59, 73, 91, 109 clones expressing egfp on, 85–86 cultures induced by, 114 promoter repression by, 21f recipes for, 139 Labeling of DNA probe by PCR using digoxigenin, 57–58 Laboratory equipment checklist, 7–8 at each lab station, 121 preparation of, 125 shared, 121 Laboratory safety, 2–3 Lac operon, 20 LacI, 20, 21f LB broth, recipe for, 126 LB/kan. See Luria-Bertani/kanamycin Ligation, 26 controls, 41–42 of DNA fragments, 40 E. coli transformation and, 39–41
electrophoresis of, reactions, 43–44 of expression vectors, 40 gel electrophoresis of, and controls, 44f transformation controls and, 41f Linear DNA, transformation of, 40 Lowry assay, 116 Luria-Bertani/kanamycin (LB/kan), 50, 63 Lysis buffer, recipe for, 134 Master mixes calculations, 83t making, 83 Master plate, preparation of, 73 Micropipettes, 7–8 calibration of, 11 definition, 9 instruction for using, 9–10 settings, 10f volume ranges of, 9t Microtiter plate, 119t Mineral oil, 58 Mini trans-blot cells, 99f Minipreps, 77 DNA alkaline lysis of, 82–83 electrophoresis of, 81 ethanol precipitation of, 82–83 isolation and characterization of, from potential transformants, 81–84 restriction enzyme analysis of, 83–84 inoculate cultures for, 77 recipes for, 137 Monoclonal antibody probes colony hybridization, 59–62 completion, 67–68 principle of, 60f Nco I, 22, 29, 40 Neutralizing solution, recipe for, 133 New England Biolabs broad range molecular weight marker, 97, 98f Nitrocellulose membrane cellular proteins binding to, 60f in SDS-PAGE and Western Blot, 98–101 Nitrocellulose spot test, 11–12 NovaBlue, 14–16 Nucleotide sequences of gfp::cbp fusion, 145f Nylon membranes, 54, 57 Oligonucleotide synthesizer, 69–70 Orientation, of expression vectors, 20–22 P20, 9 P200, 9–10
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Index PCR. See Polymerase chain reaction pEGFP-N1, 13 crystal violate stained agarose gel of, 32f egfp excising from, 29 restriction digestion of, 30 Peroxide stain, recipe for, 135–136 pET-41a (+), 21–23 preparation of, to accept egfp, 19–20 salient features of, 20f Phe64, 5 Pipette tips, 9 Plasmid DNA with cbp open reading frame, 145f containing plant promoter, 145f large scale purification alkaline lysis, 14–16 anion exchange chromatography, 14–16 DNA quantification, 17 isolating, in E. coli, 13 prep list for, 123 Polylinkers, 19 Polymerase chain reaction (PCR), 26, 54, 26, 27f, 30, 45, 53–54, 64 analysis of, 79 applications of, 70 DNA probe labeling, 57–58 gel electrophoresis and, 79 orientation of primers, 70f primer sequences, 70–71 principle behind, 69 for recombinant clones, 71–72 screen for recombinant clones, 71–72 screening, 69–72, 79 transformants by, 71f typical cycle of, 69–70 Positive clones, 49, 53, 77 fusion protein expression from, 89, 91, 95–101,103–105 replica plate, 105 Prep list, 123–142 aliquots, 124 antibiotics, 123 autoclaving, 124–125 bacterial waste, 124 E. coli host strains, 123 lab preparation, 125 plasmids, 123 recipe list, 126–127 students, 125 supplies and reagents, 125–126 Preparation of chemically competent cells by calcium chloride treatment, 35–36 Primers, synthesizing, 57–59, 60–70 Promoter repression, by LacI and IPTG, 21f Protein purification. See Purification protocols
Proteinase K, 57 Purification fractions, 115–116 analysis of, 117–121 Bradford assay of, 115–117 SDS-PAGE of EGFP::GST, 116f Purification protocols, 13 bacterial suspension cultures for, 112 inoculate cultures for, 107 Purifying protein by affinity chromatography, 113–114 QIAfilter Cartridges, 14–15, 16f Plasmid Maxi Kit, 13–14 flowchart, 16f Qiagen protocol, 13 QIAGEN® Plasmid Purification Handbook, 14 QIAquick PCR purification Kit Protocol, 26 spin, 26, 27f Reading frame, expression vectors, 21f, 22, 59 Recombinant clones, 47, 50, 65–74 characterization of, 67–73, 77, 79, 81–84 PCR for, 71–72 Recombinant protein, extraction of, from E. Coli, 107, 109–114 Removing insoluble debris from crude homogenates, 115 Replica plate positive clones, 103 Replica plating defining, 49 petri plate grids for, 51f positive clones, 50, 51f, 105 process for, 50 Repression, 21f Restriction analysis of miniprep DNA, 83–84 Restriction digestion of pEGFP-N1, 30 of vector restriction enzyme digestions, 23 Restriction enzyme analysis, of miniprep DNA, 81–84 RNase, 13, 83, 83t Safety, 2, 54 Sample buffer, recipe for, 24, 97, 140 Screenable marker, 19 Screening transformants, 45 counting, 49 by PCR, 71f SDS-PAGE. See Sodium dodecyl sulfatepolyacrylamide gel electrophoresis Selectable marker, 19–20 Ser65, 5, 29 Serial dilution scheme, 11t
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Index Silver stain, 96 SOC broth, recipe for, 131 Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), 95–100 of EGFP::GST purification fractions, 101–103, 115–120 inoculate cultures for, 93 purification fraction analysis by, 115–120 of purified fusion protein, 117 Western Blot and apparatus preparation in, 96–97 blocking, 100 electrophoresis, 98 incubation of blot with antibody against GFP, 101–102 incubation of GAMP, 102–103 nitrocellulose, 101, 102f preparing molecular weight standards, 97 sample loading, 97 sample preparation, 97 Southern Blots, 45 Spin column, cloning DNA using, 26–27 SSC, recipes for, 133, 135 Station checklist, 7–8 Stringency, 53–54 washes, 64, 111 Students, preparation of, 125 Support matrix, 110, 111, 111f T7 promoters, 20, 35, 73 TBE stock, recipe for, 127 TBS. See Tris-buffered saline TE buffer, recipe for, 126 Thermus aquaticus, 69 Thr, 5, 29 Transfer buffer, recipe for, 140 Transformation and storage solution (TSS), recipe for, 130 Transformation competent cells
calcium chloride treatment in, 35–36 control, 36–37 E. coli, 35 freezing, 36–37 control, 36–37 DNA ligation and, 41f divalent cation-mediated, 42–43 miniprep DNA, 81–84 of E.coli, DNA ligation and, 39–41 Tris-buffered saline (TBS), recipe for, 140 Tris-glycine running buffer, recipe for, 139 TSS. See Transformation and storage solution Vector restriction enzyme digestions, restriction digestion of, 23 Vectors. See also Cloning vectors; Expression vectors in ligation reactions, 43 Visualization of clones expressing egfp on IPTG, 85–86 Volume ranges, of micropipettes, 9t Wash buffers, recipe for, 134 Wash solutions, recipes for, 135 Water, density of, 11 Western Blot, 45, 91, 95 experimental flowchart, 102f SDS-PAGE and apparatus preparation in, 96–97 blocking, 100 electrophoresis, 98 incubation of blot with antibody against GFP, 101–102 incubation of GAMP, 102–103 nitrocellulose, 101, 102f preparing molecular weight standards, 97 sample loading, 97 sample preparation, 97 YT broth, recipe for, 139
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Emergency Contact Form Fill out the following form (or copy of the form) and turn it in to your instructor. The information on this form will be used in case of emergency.
Name
Laboratory section number and day
Person to contact in case of emergency
Telephone number(s) of emergency contact
Drug allergies and other relevant medical conditions
I have received and understand the safety training for this laboratory course. Signed: Date:
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