Fall 2003: GENE EXPRESSION
ANSC/GENE 626 (edited by. N. Ing 9/2/03)
Nancy H. Ing, Instructor
Kleberg 410D, 862-2790
ning@cvm.tamu.edu

The purpose of this course is to provide graduate students with experience in working with RNA and DNA and with the theories behind the use of molecular biology in research.

Class will be held in BICH 243 in Fall semester on Thursdays (lecture from 12;40 to 1;35 p.m.) and Fridays (lab from 12:45 to 3:40 p.m. )

13. Plasmid DNA restriction and gel analysis

Protocols will be provided. Required viewing of videos. Required reading from Gerstein "Molecular Biology Problem Solver" as well as other materials relating to kits, etc used. Students will be evaluated on preparation and participation (33%), lab notebook (33%) and a written laboratory report (33%) due the Friday after the last class.

ANSC/GENE 626 REQUIRED READING & VIDEOS

• Required Text: Gerstein "Molecular Biology Problem Solver" 2001
  
• Lab 1 Pipettors pp 67-77.
  
• Lab 2 RNA purification pp 197-224
  
• TriPure handout - see Roche website at http://biochem.boehringer-mannheim.com
  
• Isolation of mRNA Video*
  
• Lab 3 SPectrophotometers pp 94-111
  
• Ambion Maxiscript and Northernmax (formaldehyde gel) Protocols - see ambion website at www.ambion.com
  
• Lab 4 Gel Electrophoresis pp331-371
  
• Polyacrylamide Gel Electrophoresis Video
  
• Lab 5 Nucleic acid hybridization pp 399-460
  
• Lab 6 DIG RNA labeling kit instructions from Roche (see website biochem.roche.com)
  
• Southern Blot Hybridization Video
  
• Lab 7 DIG luminescent detection kit from Roche
  
• Lab 8 Clontech MIMIC kit information
  
• Polymerase Chain Reaction pp291-330
  
• PCR and Detection Video
  
• Lab 9 Agarose Gel Electrophoresis Video
  
• Lab 10 DNA Ligation Video
  
• Lab 11 Transformation Video
  
• Lab 12 DNA Purification pp167-195
  
• Rapid Isolation of Plasmid DNA (minipreps) Video
  
• Lab 13 Restriction enyzymes pp225-266
  

*Videos may be checked out from Kleberg 410A for 1 h during the day or overnight after 4PM (returned by 9AM). Single and group viewing is available in the Multi Media Center (Kleberg 023).

LAB RULES

A. GENERAL

1. Everyone is individually responsible for the experiments. Come prepared by reading protocols and required reading in advance!! Activities will be started immediately, while explanations and discussion sessions will occur as time permits. COME PREPARED to ASK QUESTIONS, especially during discussions.

2. Equipment in this and neighboring labs is shared. Know or ask how to use it. Obey user rules, such as signing logs. Leave all equipment in good working order. If there are problems, tell someone so we can fix them!

3. Leave the lab better than you found it. Wash your own glassware, clean up your work area, write the names of reagents that are running out on the "to be ordered" list, etc.

4. Lab notebooks are bound volumes, are kept in pen with numbered, dated pages. They are designated only to that purpose, are labelled on the cover with Name, dates and lab, and are only removed from the lab with instructor permission. In them, each person describes their activities and observations each day. Each page should be dated. Each photo or X-ray film should be labeled with initials, date, and identification of gel type, lane contents and dye migration. Record things chronologically along with dates. Start a new project on a new page with a description of its purpose. A stranger should be able to pick up your notebook and understand how and why you did an experiment. Protocols don't have to be written each time, but may be referred to, instead. For this, you may consider the handouts you receive in lab as references. Use them as such and refer to specific pages (dates) within them. Record your activities and observations clearly, using complete and understandable sentences. Write down your reagents, including buffers and buffer recipes. KEEP UP WITH NOTEBOOK ENTRIES EVERY DAY…otherwise, data will be lost!

5. All reagents and samples saved must be labeled with the date; your initials, and WHAT IT IS. In my lab, we use simple sample numbers, such as 1 - 10. To key it into our notebook, we write: lab notebook #, page #; (NI2P30 relates to Nancy Ing Book 2, page 30), so the notebook, not the tube, is where the full description of the sample exists. Items not labeled sufficiently may be discarded.

6. Store things in appropriate places! For plasmids and reactions and buffers, store at –20C in storage box provided to your group unless otherwise noted. Note storage places in your notebook.

B. SAFETY IS THE #1 PRIORITY

1. The only safety activity not strictly enforced is wearing safety glasses: this is a good idea but is not mandatory. Wearing a lab coat is mandatory and wearing gloves will become a habit (see below).

2. Working with open flames and hazardous chemicals have strict safety protocols - ask for them and follow them.

3. Working with radioactivity is a privilege, not a right. Workers must monitor for contamination, before, during, and after the procedure. Radiation safety training is required. WE RUN A CLEAN LAB.

4. Because we work with HAZARDOUS SUBSTANCES, there is NO EATING, DRINKING, SMOKING, or APPLYING MAKE-UP in the lab.

5. Garbage must be disposed of properly. Glass and sharps, biohazard, chemical, and radioactive waste must be separated from the rest.

C. GOOD LAB TECHNIQUES

1. ICE IS NICE! Work on it unless otherwise directed. It slows degradation of macromolecules.

2. Many reagents settle on storage, so mix them! All frozen solutions need to be thawed and mixed before using.

3. ENZYMES DO OUR WORK. They are stable as glycerol solutions at -20?C. Keep them in the freezer as much as possible. Only remove them in -20oC blocks. DO NOT WARM ENZYME STOCKS! When pipetting small amounts of viscous solutions like enzymes, check loaded pipet tip and evacuated one to assure that enzyme got into the reaction. After addition, mix reaction solution gently but thoroughly: can pipet total volume up and down OR vortex gently and flash spin in microfuge to return reaction to the tube bottom.

ADA Statement, Copyrights, and Plagiarism

The Americans with Disabilities Act (ADA) is a federal antidiscrimination statute that provides comprehensive civil rights protection for persons with disabilities. Among other things this legislation requires that all students with disabilities be guaranteed a learning environment that provides for reasonable accomodation of their disabilities. If you believe you have a disability requiring an accomodation, please contact the Dept. of Student Life, Services for Students with Disabilities in Room 126 of the Koldus Bldg. or call 845-1637.

Copyrights

The handouts used in this course are copyrighted. By handouts", I mean all materials generated for this class, which include but are not limited to syllabi, quizzes, exams, lab problems, in-class materials, review sheets, and additional problem sets. Because these materials are copyrighted, you do not have the right to copy the handouts unless I expressly grant permission.

Plagiarism

As commonly defined, plagiarism consists of passing off as one’s own ideas, words, writings, etc., which belong to another. In accordance with this definition, you are committing plagiarism if you copy the work of another person and turn it in as you own, even if you should have the permission of that person. Plagiarism is one of the worst academic sins, for the plagiarist destroys the trust among colleagues, without which research cannot safely be communicated.

If you have any questions regarding plagiarism, please consult the latest issue of the Texas A&M University Student Rules, under the section "Scholastic Dishonesty."

BICH/GENE 432

Safety Precautions

1. ABSOLUTELY no food or drink in the lab. This includes gum.

2. Always wear gloves when working with Ethidium bromide, radioactive compounds, acrylamide and organic compounds.

3. Always wear UV safety glasses when using UV illumination.

4. Use special care when working with open flames. Don't forget to turn off gas after use.

5. Clean up spills immediately. Notify instructor if hazardous compounds are spilled.

6. Discard organic solutions in appropriate bottles.

7. Discard Ethidium bromide waste in correct container.

8. All culture medium and labware used for bacteria needs to be autoclaved or put in Chlorox before disposal or washing.

9. After using radioactive materials, wash the work area and clean spills immediately. Dispose of gloves in radioactive waste as soon as you are finished.

10. All sharps (including broken glass, needles and razor blades) should be disposed in clearly marked containers, not in the general trash.

11. If you have questions about anything, ASK!

12. Lab coats are required!

LAB REPORTS - BICH/GENE 432
by Linda Guarino

TITLE - This is the most important part of a manuscript. A reader begins here, and will also finish here if the title does not promise a subject of interest to him/her. A good overall rule is to use the fewest possible words that adequately describe the contents of the paper. But, do not sacrifice words for specific information. For example 'DNA cloning' is a short title, but it is too general. A popular trend in recent years is to publish papers where the title is a complete sentence that summarizes the major conclusion of the manuscript. Personally, I prefer titles that describe the work, not the results.

ABSTRACT - An abstract is a mini version of the paper. It should provide a brief (less than 250 words) summary of the major points of the manuscript. The abstract should state the objectives, describe the methodology used, summarize the results, and state the principle conclusions. The abstract should be written in the past tense, because it refers to work done.

INTRODUCTION - First of all, state the nature and scope of the problem investigated. Review the pertinent literature(NOT NECESSARY FOR THIS CLASS). Describe the method of the investigation. State the principle results. State the principle conclusions suggested by the results. The first two parts should be in present tense, while comments relating to the present study should be in past tense.

METHODS - The methods section should expand upon the description of the methodology that was presented in the abstract. The order of presentation is usually chronological (methods used in initial stages of the study are presented first). However, sometimes it makes more sense to group similar methods into sections, even though they were not used at the same time. Due to space limitations in journals, methods are not usually described in detail if they have previously been published. If a scientist uses a protocol that is identical to one previously described, he/she would state 'The DNA was prepared according the procedure previously described (reference). If there were minor differences, he/she would state 'according to the procedure of (ref.) with minor modifications' and then describe the modifications. In this class, you may assume that the class protocol has been published. Therefore you don't need to give the details, but you need to describe the general strategy. For example, you should say 'the DNA was purified by the alkaline lysis procedure as previously described' not 'the DNA was purified as previously described'. In addition to the class protocol, you could also reference the Cloning manual or the Promega manual. The methods section should be written in past tense.

RESULTS - The results section is a presentation of the data. It should not repeat the methods given in the previous section. Each figure should be referred to here. The results section should be written in past tense.

DISCUSSION - The discussion should put the results into perspective. Discuss the results without recapitulating the results section. Show how your results and interpretations agree. State your conclusions clearly, and summarize the evidence for each conclusion. Selection of correct tense is more difficult in the discussion than in the other sections. Your own work should be described in past tense. If reference is made to published work, it should be in present tense.

REFERENCES - Only need to cite the class protocols and any other sources of material...no need for literature review so these are very few.

FIGURES and FIGURE LEGENDS - Present the important data in figure form, raw data if possible for this class. Figure legends begin with a title for the figure. Figures should have complete legends - so that they can be understood without reading the rest of the paper. These may be nested in the paper or placed at the back.

Many students have asked about length. The best rule that I can give you is that it should be long enough to convince me that you have learned something. However, I have a short attention span, and if your paper is very long and verbose, I may lose interest before I decide whether or not you have learned anything.

Suggested reading:

Day, R. A. 1988 How to write and publish a scientific paper, 3rd ed. Oryx Press, Phoenix.

9/6/02

Introduction to Gene Expression

General

The purpose of this class is to provide motivated students with the beginning skills required to apply molecular biology techniques. Protocols and hands-on exercises will teach several techniques, but more importantly, will allow new procedures to be mastered during subsequent research projects. Students will learn how to handle nucleic acids through basic purification and measuring and manipulative procedures. However, comprehensive coverage of the underlying biochemistry of DNA modulatory enzymes, for example, is not possible in the time allotted. Excellent courses exist for such background studies (GENE 431 and 450), and required reading and suggested references will fill those voids. Brief discussions of how applications of molecular biology are used in physiological studies will be held as time permits. Questions are encouraged, also. We will have free time during incubation periods - use these wisely with questions, reading or writing for your notebook or lab report.

The major criticism of this course is that the flow of experiments overlaps. This is life. Experiments proceed too slowly with long incubation times to just sit and wait. It's up to us to keep the purpose of the experiments in our minds (and notebooks!).

This course is designed to help new students study expression of their favorite gene in their favorite tissue. For this class, your favorite genes are actin and glyceraldehyde phosphate dehydrogenase (GAPDH), because they are highly expressed in your favorite tissue (endometrium.) Genes are transcribed into messenger RNAs, so first we'll:

1) Extract RNA from tissue and use it to make a Northern blot. The amount of RNA obtained is measured by sample absorbance at 260mm. To assess RNA quality, the RNA preparation is analyzed on a denaturing gel. This is transferred to a membrane ("Northern blotting") for hybridization with probes for specific mRNAs (actin or GAPDH)..

2) Make a cRNA probe for actin or GAPDH mRNA and hybridize it to the tissue RNA on the Northern blot. The lab boss gives out plasmid clones containing complementary DNAs (cDNA's, synthetic copies of fragments of mRNAs). The best probes are cRNAs. To make them, the circular plasmid DNA is restricted or cut at a specific site with a restriction enzyme. For in vitro transcription, the plasmid and ribonucleotides are combined with a bacteriophage RNA polymerase (SP6 or T7). The polymerase enzyme binds a specific site on the plasmid and transcribes (makes RNA) using the DNA as a template. Either of the two strands of the DNA can be reproduced as cRNA: the top cDNA strand is like mRNA and, if transcribed, is called "sense" cRNA. The sense cRNA is useful as a template for translation. The bottom cDNA strand is complementary to the sense strand and hybridizes to mRNA, as does its transcription product called "antisense" cRNA. We'll only synthesize the antisense cRNA and it will be labeled with Digoxigenin so it will be a probe (detectable reagent) for identifying its homologous RNA in the tissue RNA samples. After hybridizing the probe to the RNA on the Northern blot and washing the blot, specifically bound probe will be detected with DIG-antibody and chemiluminescent detection reagents.

3) Quantitate estrogen receptor mRNA with RT-PCR. But what if you only have a very small amount of tissue and/or the mRNA you want to study is rare (such as estrogen receptor (ER))? The most sensitive mRNA quantitative technique is quantitative Reverse Transcriptase-polymerase chain reaction (PCR). Reverse transcriptase copies RNA into cDNA. The reverse transcription of tissue RNA will provide cDNA that will be used to amplify a specific target cDNA between two previously designed primers Since PCR is fickle, the best way to make it quantitative is to use an internal control DNA that competes for the same primers and reagents but can be distinguished (by slightly different size on agarose gels). Therefore, one runs the PCR reactions on an agarose gel and simply looks for the lane with equivalent bands (PCR products) from the internal control and target cDNAs in a titration set of PCR reactions. Since product amounts are the same, starting material must be the same; thus mRNA concentration = internal control concentration, the latter of which is known for that reaction.

4) Subclone beta-globin cDNA to a more useful plasmid vector. But what if the cDNA clone you have is in a plasmid vector that doesn't have SP6 or T7 RNA polymerase sites (many old plasmids don't). You may have to sub-clone the cDNA fragment into your desired vector. The easiest way utilizes PCR to amplify the cDNA. Restriction enzyme sites can be created by synthesizing them on the ends of the primer. This allows easy insertion and ligation of the cDNA and vector. The new plasmid is forced into E. coli cells during transformation, where presence of the plasmid confers a new phenotype to the bacteria: resistance to the antibiotic ampicillin. Clonal colonies are grown and their plasmid DNA prepared (mini-preps) individually to identify the desired clone (by restriction enzyme analysis).

Thus, our different lab exercises all fit together into common techniques utilized in studies of gene expression. KEEP THE FLOW OF LOGIC IN MIND! Don't just come in and mix reagents and shuffle tubes. Think of the molecules and what you want to learn from them. Predict (visualize) the results of your experiments before you perform them.

YOUR SUCCESS DEPENDS ON PREPARATION:

You must read and think through the experiments BEFORE THE LAB to be able to perform and interpret them well. Note that grades are 33% preparation!

Lab 1 9/6/02

Preparing Materials for RNA Work

Beating RNase

You need to read about RNase, a ubiquitous enzyme that efficiently destroys RNA. Primarily, this will serve to make you paranoid and do neurotic things, like wear gloves all the time. Although working with RNA is similar to working with DNA, many RNA experiments fail miserably because of RNase, so know this enemy!

In biochemistry, RNase is the model of an enzyme that will not die: not in an autoclave or even after dehydration (by alcohol, etc.). As soon as it returns to a water environment between room temperature (R.T.) and 37oC, it chews again. It is an enzyme of all living things and is important in keeping RNA turnover high so cells don't choke on RNA and so new expression of genes tightly regulates cell function.

The best way to beat RNase is to avoid it. Work with the cleanest reagents and lab-ware. Things that aren't handled by living thing are generally RNase-free; e.g. paper towels. Test tubes and pipettes don't have to be sterile but should be used from freshly opened packages. Then protect packages from dust and fingers by resealing packages and storing in cabinets. Glassware is reserved similarly: wrapped and stored away from general use. Equipment like Pipettemen and Gel apparati for RNA are reserved for this use and are NOT USED WITH RNase!

Solutions are made with water of the highest purity. Dry chemicals are shaken out of containers: residual amounts are discarded. Nothing dirty is introduced into chemical stocks: solutions or powders.

All solutions are treated with 0.1% diethyl pyrocarbonate (DEPC). This oily liquid is added. The solution is shaken vigorously until foamy (aerobic workout). The solution is incubated 37oC overnight to allow the DEPC to covalently attack RNase. The solution is then autoclaved to destroy DEPC (which also attacks RNA) and to sterilize to prevent growth of undesirables. Exceptions to this solution preparation protocol are 20% SDS (nothing grows in this) and Tris solutions (which DEPC attacks, too). NOTE: DEPC treatment can only correct a low level of RNase contamination! You must start clean!!!

Today's Exercise

Apply benchcote and work with gloves on and tubes/racks on top of diapers.

A. Pack tips: 1 blue box and 2 yellow boxes per group

1. Pour clean blue tips onto a clean surface (paper towel or diaper).

2. With new gloves, pack tips.

3. Write name on box & protect it!

4. Repeat with yellow tips: fill 2 boxes.

B. Practice pipetting

1. Read instructions for pipetting in Appendix.

2. Tare a 1.5 ml test tube on the balance.

4. Weigh 1000 ul of water two times.

5. Repeat with Isopropanol.

6. Determine the densities of these liquids.

7. Measure the volumes of the unknown samples provided in the 1.5ml test tubes.

8. Check your results with an instructor.

C. Each person should test tap water, distilled and DEPC-treated water for Rnase:

1. Label 4 Rnase-free 1.5 ml tubes. Pipet 4 ul of either tap, distilled or DEPC H2O into each (two tubes get DEPC-H2O).

2. Pipet 1ul test RNA into each tube.

3. Put all but one DEPC-H2O tube in 37oC block for 1 h.(The lone DEPC-H2O tube should be kept on ice during the 1 h incubation.)

4. Store all of the tubes at -80oC until Lab 4.

D. Make 1 li DEPC-H2O and 1 li of 20XSSC per person

1. In 1 liter bottle add nanopure H2O to 1 liter level for DEPC-H2O. For 20X SSC, add 175 g NaCl and 88 g Na citrate to a 1 li bottle and dissolve in nanopure H2O.

2. Add 1ml DEPC per liter.

3. Shake till foamy for 10 sec.

4. Put in 37oC incubator O/N to allow DEPC to work optimally.

Lab 2 9/13/02

RNA Extraction

[Autoclave the DEPC-H2O to destroy DEPC and prevent any growth in solutions that might introduce RNase.NOTE on Autoclaving: Need 35 to 40 min. sterilization time for 1 liter. 20 min. for 500ml. Use "liquid" cycle and keep caps loose]

You know how to fight RNase to keep materials clean. GUESS WHAT! RNase is in all living systems including the one from which you'll purify RNA. So all RNA preparers begin with the realization that their worst enzyme enemy is present in the sample. In the cell, RNA is compartmentalized away from RNase so many tissues are OK for harvesting for RNA if kept cool 2-6 h after collection (of course, faster may be better). But freezing breaks intra-cellular membranes, mixing RNA with RNase. Therefore, fresh tissues are kept cool while mincing and weighing, then are put in a 1.5 ml polypropylene tube snap frozen in liquid N2. They are stored at -80?C. They may store well for 6 months but usually not for 1-2 years. This is dependent on them never thawing, too. So the TWO MAIN POINTS about tissue collection are to SNAP FREEZE and KEEP at -80oC until use within 1 year. NOTE: You can't snap freeze things much bigger than 0.5 cm3. I mince to about 5 mm or less. Tissues vary with RNase content and amount of connective tissue present, so RNA yields vary in quality and amount. RNA extraction from cultured cells results in very high quality RNA, usually.

RNA Extraction from tissue with Boehringer Mannheim TriPure reagent (contains phenol! see NOTE 1 below)

Each student will do 2 RNA preps (one from endometrium, one from spleen). Label all tubes needed NOW!

1. Homogenize 0.5 mg tissue (frozen or fresh) in 5 ml room temperature ("RT") Tripure solution in a 50 ml polypropylene tube. Use three 15 sec bursts at 70% power. Rinse probe in tripure (do a mock homogenization with Tripure and no tissue) between dissimilar samples.

2. Incubate RT 5min. During this time, transfer the contents equally into 4 - 1.5 ml tubes.

3. Add 250 ul chloroform using a P-1000. Mix by vortexing or shaking vigorously 15 sec.

4. Incubate RT 5 min.

5. Centrifuge 15 min at 10,000 rpm at RT or 4oC in a microfuge.

6. Transfer upper phase to four clean 1.5 ml tubes with transfer pipet. AVOID THE INTERFACE!!!! Discard lower phase and interface in phenol waste container.

7. Precipitate RNA by adding an equal volume of isopropanol. Mix by inverting tube. Incubate RT 5 min

8. Centrifuge at 10,000 rpm for 10 min at RT.

9. Wash pellet in 75% EtOH (make 10 ml with 100% EtOH and DEPC H20). This means to discard the supernatant, add the supernatant volume of wash (75% EtOH), vortex, microfuge 5 min, and discard supernatant. The purpose is to wash salts out of the RNA pellet, which should not dissolve during the procedure.

10. Air dry pellet briefly after spin. You can wipe the sides of the tubes with Kimwipes, but stay away from the pellets! Do not dry totally or you will not be able to solubilize RNA easily!!!

11. Store pellet at -80oC.

NOTE 1: Tripure has phenol and guanidine salts in it...both are caustic and burn skin!!! Be careful! Wear safety glasses!!!

NOTE 2: CHCl3 (Chloroform) dissolves things like styrofoam and polystyrene - use glass graduated pipets and polypropylene 15 & 50 ml tubes.

BEFORE YOU LEAVE, CLEAN UP AND RECORD ACTIVITIES IN NOTEBOOK!!!

NOTE 3: Record observations in notebook!

Examples:1) Lysate in step 2 was viscous!

2) Tube #2 fell and was lost.

3) RNA pellet #5 took 20 min. to dissolve, while #1 took only 1 min.

Lab 3 9/20

Analyzing Extracted RNA by Absorbances; In Vitro Transcription of cRNA Probes

A. Solubilize RNA samples from tissue and measure Absorbance at 260 nm and 280 nm

Absorbance measures of DNA & RNA at 260 nm are used to estimate concentrations of nucleic acids. An Absorbance of 1.0 for solution of double-stranded (ds) DNA has =50 ug/ml while RNA has A260 =40 ug/ml and single-stranded (ss) DNA A260 =37 ug/ml. An unknown sample of RNA can be measured for A260 and [RNA] = A260 X dilution factor X 40

The ratio of A260/A280 is an indication of the purity of the nucleic acid. The ratio for pure aqueous DNA is 1.8 while for RNA it is 2.0. Protein, phenol, EtOH and other things often lower these ratios because they absorb at A280.

1. Dissolve the four similar pellets each in 25 ul 1 mM Na citrate Buffer/pH 6.4 or TE buffer (10 mM Tris, 1 mM EDTA pH8).

(Heat in 70oC block and vortex hard and repeatedly over 15 minutes.) Pool so that you have a 100 ul sample for each RNA prep.

{RNA STORAGE: Store at 4oC during sample use (this class). For storage over 1 week, can store at -80oC. For longer storage, add 3 volumes of ethanol and store at -80oC.}

2. Add 0.5 ml of DEPC-H2O to seven 1.5 ml tubes.

3. Label one "Blank" and the others (duplicates) after the samples: the two RNA samples and a positive control: 10mg/ml salmon sperm DNA.

4. Add 2 ul aliquots of samples to @ tube except "Blank".

5. Use micro UV-transparent disposable cuvettes and rubber or plastic transfer pipettes. Blank the machine to read 0 absorbance at A 260 nm with the "Blank" in a cuvette. Then repeat with dilute samples in cuvettes. Repeat the procedure for measuring the A280 of samples.

6. Estimate [RNA] (ug/ml) = A260* diln. Factor* 40

= A260 * 250 * 40

Therefore: [RNA] (mg/ml) = A260 * 10 (= ug/ul, too!)

7. Pipet 32 ug RNA into a clean 1.5 ml tube for each RNA prep. We want 32 ug aliquots of RNA for running replicate 8 ug samples on a Northern gel. If volume is less than or equal to 15 ul, store "as is" at -80oC. If volume is greater than 15 ul, precipitate the RNA by adding 3 volumes of 100% EtOH and 0.1 volumes of 3 M NaAc/pH 5.2. Vortex and store at -80oC.

8. To the rest of the RNA preps, add 3 vol 100% EtOH and store at -80oC. (This is a good way to store RNA without degradation for years.)

B. In vitro transcription - DO THIS FIRST!

NOTE : There are 3 common types of nucleotide probes: DNA oligonucleotides (ss), cDNA (ds) and cRNA (ss). For many applications, cRNA probes are superior over:

1. end-labelled oligonucleotide probes because they are:

a. longer (and therefore carry more label and have higher hybridization specificity)

b. uniformly labelled throughout (so they carry more label)

2. nick-translated or random-primed cDNA probes, because they only have the desired probe strand, not the other "sense" strand that increases background.

In addition, the binding of RNA:RNA hybrids is stronger than that of DNA:DNA hybrids.

REMEMBER:

1.the cDNAs are synthetic cloned fragments of the mRNAs

and

2. Knowing the information on the plasmid maps (See appendix) is critical to designing the probes (e.g. knowing what enzyme to linearize the plasmid with, which enzyme to transcribe with, and which strand (sense or antisense) is generated.

Circular plasmids must be linearized with a restriction enzyme to generate DNA templates suitable for in vitro transcription. You will use SP6 or T7 RNA polymerase to transcribe antisense cRNAs for actin and GAPDH mRNAs and 18S rRNA. Ambion's "pTRI-____" constructs are foolproof: already linearized, and have all the RNA polymerase sites on the side of the cDNA so as to make only antisense transcipts. (REMEMBER THAT RNA IS SINGLE-STRANDED!!!)

ANSWER THESE and all other PROTOCOL QUESTIONS IN YOUR NOTEBOOK

Q1.: If we wanted to probe a Northern blot with cRNA from poPR77A, which restriction enzyme and RNA polymerase would we use with that plasmid? (see map).

Q2.: If we wanted to do in vitro translation with cRNA from poPR77A, which restriction enzyme and RNA polymerase would we use?

[In vitro transcription kits can be obtained from various sources. Ambion's "Maxiscript" is OK and is the basis for the following reaction (one per group).]

A. In vitro Transcription using the Roche DIG-Labeling kit.

Each student should set up one in vitro transcription reaction for DIGOXIGENIN-labeling antisense 18S, GAPDH, or actin cRNA probes:

1. Thaw components at room temperature (RT) then store on ice.

EXCEPTION: RNasin and RNA Polymerase, like all enzymes, stay at -20oC always!

2. For sense cRNA add components from kit, in order, to a 0.5 ml tube at RT.

12 ul DEPC-H2O

2 ul 10X Transcription Buffer

1 ul RNasin

2ul 10 mM rATP, rCTP, rGTP, and 3.5 mM DIG-11-UTP

2 ul linearized DNA template

1 ul SP6 or T7 RNA Polymerase

3. Mix by pipetting up and down...gently! No bubbles.

4. Flash spin in a microfuge

5. Incubate 37oC for 1h

6. Add 1 ul RNase-free DNAse

7. Incubate 15 min at 37oC

8. Save 5 ul of each of the transcription reactions in 1.5 ml tubes for analysis on a urea 5% acrylamide short fat sequencing gel. Store these tubes and the original reaction tubes at -80oC.

9. For short fat acrylamide gels in next lab, wash 1 short and 1 long plate with soap and water and a 250 ml Erlenmeyer flask or beaker. Soak in 1 M NaOH overnight (O/N).

Lab 4 9/27

In Vitro Transcribed RNA (and RNase test) analysis on urea/acrylamide gel

A. Analyze in vitro transcription products on a 8M urea, 5% acrylamide gel called a "probe test gel or short, fat sequencing gel." These denaturing acrylamide gels are especially suitable for analysis of small (<400 base) DNAs and RNAs.

1. Make a probe test gel. (See Appendix recipe and plate setup for vertical gel.)

2. Add 15 ul of deionized formamide loading dye to your 5 ul cRNA aliquots as well as to your RNA samples from the first lab (RNase test). Similarly, prepare a sample of RNA Century (Ambion) markers. Leave the 15 ul aliquots of the cRNA reactions at -80oC!

3. Heat 68oC for 5 min.

4. Cool on ice.

5. Flush wells free of urea with 1X TBE in needle & syringe.

6. Load with elongated gel loading tips carefully! in the bottom of wells.

7. Run at 35 mamps for 1 h.*

8. Disassemble gel plates.

9. Stain the gel with ethidium bromide by soaking in 2 ug/ml EtBr for 5 min. Use gentle agitation.

10. Destain by soaking in d H2O for 5 min. with agitation.

11. Photograph the gel on a UV light box. Label the photograph completely (see Lab Rules A4) and tape it in your notebook.

12. Compare migration distances and brightnesses of bands with those in RNA marker lane. Note the migration of the tracking dyes, too (see table in APPENDIX) (Xylene cyanol is light blue & slow, while bromphenol blue is dark blue and fast).Mark the dye migration positions on the photograph so you could rerun the gel exactly if you wanted to.

13. For the next lab, treat a 250 ml Erlenmeyer flask or beaker and a mid-size horizontal gel apparatus with 1 M NaOH. Leave these to soak until next time.

14. Do calculations for Lab 5 step A6. Dilute a small amount of plasmid DNA template for in vitro transcription to 10 ng/ul.Store at -20oC.

*During the gel run, discuss DNA structure using human nucleotide models:(answer all questions in your notebook!)

a. Make a single-stranded six base random sequence and identify 5' and 3' ends. What types of bonds exist between the bases?

b. What is the chance that a specific 6 base sequence will occur randomly?

c. Reverse the sequence polarity. What bonds did you have to break?

d. Make a complementary strand of DNA. How are the strands oriented to each other? How is this strand's sequence related to the initial strand? What types of bonds are between the strands?

e. Make an EcoRI restriction enzyme site. What bonds does the enzyme cut? What makes the ends "sticky"? Do the ends have 5' or 3' overhangs?

Lab 5 10/4/02

Northern Gels (NorthernMax (Ambion) protocol) and Blotting

Be clean - protect apparati from Rnase contamination- clean diapers!

All DEPC reagents

A. Recover 32 ug total cellular RNA samples from the endometrial and liver extracts that you made (2 per student).

For each RNA prep, make a 32 ug sample to provide duplicate samples for two lanes. If samples from C7 of Lab 3 are precipitated start at 1 and continue. If not, start at step 6.

1. Spin 10 min at 4oC at 10,000X g

2. Discard supernatant by decanting

3. Dry sides of tubes with Kimwipe and remove residual ethanol from pellet with yellow tip if needed.

4. Air dry pellets 5 min.

5. Dissolve pellet in 15 ul 1 mM Na citrate at 68oC with vortexing.

6. If your sample was not precipitated, bring it up to 15 ul with 1 mM Na citrate. Add 45 ul Northern Sample Loading Dye and 6 ul of 0.1 mg/ml EtBr to each sample. These will be loaded in two lanes as described below. ALSO, prepare 10 ng samples of linearized plasmids with cDNAs for 18S rRNA, GAPDH, and actin (diluted to 10 ng/ul in Lab 4 step 14) in 5 ul 1 mM Na citrate + 15 ul dye. Add 2 ul of 0.1 mg/ml EtBr to each sample. Lastly, prepare a 15 ul sample of mouse liver control RNA to be split into two lanes during loading.

Q: What is the concentration of the control RNA? How much is in each lane?

7. Heat 68oC for 10 min.

8. Chill on ice

B. Prepare & Run gel. 2 people per gel: one student load the top row, one the bottom row of wells

START THIS FIRST OR SIMULTANEOUSLY WITH YOUR RNA SAMPLE PREPS (above)!

1. Rinse gel rig and beaker/flask well with house-distilled water. Melt agarose (0.8 g) in 72 ml DEPC-H2O in an RNASE-Free glass bottle or beaker. (Bring to a boil in microwave oven and mix by swirling: repeat 2 to 3 times).

2. Cool to 70oC.

3. In a fume hood, add 8 ml 10X Denaturing Gel buffer (formaldehyde and MOPS/pH 7.0, NaAc, and EDTA) and pour into RNase-free gel mold with the ends taped. Use two thin combs.

4. Load samples onto the gel under 1X Gel Running Buffer (dilute the 10X stock from the kit with your DEPC-H2O bottle, cover the gel with buffer to about 0.5 cm depth). LOADING ORDER[Skip spaces = "X"]

Millenium RNA markers, Positive control RNA , student1 Endometrial RNA, student1 Liver RNA, pTRI- actin, pTRI-GAPDH, pTRI-18S rRNA, student2 Endometrial RNA, student2 Liver RNA plasmid.

5. Run at 100 volts until dye front reaches bottom or sufficient separation occurs (1 to 1.5 h); can peek at gel progress with hand-held short wave UV lamp. Prepare materials for blotting during this time!!!!!!!!!!!!

6. Take photo on UV box alongside a fluorescent ruler.

NOTE: MARK DYE POSITIONS ON GEL PHOTOGRAPHS. Label all photos completely! Name, date, identity!!! Number lanes on photo then desribe them in your lab notebook.

C. Northern transfer to nylon membrane

1. Cut wicks, blotting papers, and nylon membrane wet in transfer buffer as directed.

2. Soak gel in 10X SSC for 15 minutes with gentle agitation.

3. Assemble an upward capillary transfer as instructed in the diagram. You can use the rig you ran the gel in to do the transfer to nylon. Allow the transfer to continue with 500 ml of 10X SSC until the next lab.

Lab 6 10/11

Northern Blot hybridization

A. UV cross-link RNA to Northern blot

1. Remove all papers from the Northern transfer but keep the gel and blot together. , mark well positions on the blot with a sharp pencil or a black Sharpie marker. Also write initials and date. Put these marks on filter's back. Keep the blot RNA-side-up during subsequent handling.

3. Rinse filter for 30 sec. in 2X SSC with vigorous agitation.

4. Put blot on plastic wrap and on UV box. On the side of the blot, mark positions of 28S and 18S rRNAs and RNA markers. Also, mark places you will cut the blot in the future (to hybridize with different probes), e.g., on the blank lane #6. You can take a picture to confirm that the RNA transfer was good. Look at the gel on the UV box. Did all of the RNA transfer out?

5. Place wet blot on top of Whatman paper saturated with 2XSSC - all on top of plastic wrap.

6. UV crosslink RNA to nylon (use Stratalinker in energy mode: 120,000 (or 1,200,000 ?) ujoules).

7. Cut the blot into pieces to be probed with 18S rRNA and GAPDH and actin probes.

B. Prehybridization & Hybridization

Since the nylon membrane likes to bind things, background sites are blocked (bound) with non-specific DNA and protein. Usually, sheared salmon sperm DNA is used in prehybe to block these sites.

1. Warm the Ambion Ultrahybe hybridization solution to 65oC. Swirl to dissolve precipitates.

2. Make three plastic bags from a tube with the heat sealer. Double seals are a good idea. The bags should be 2 cm longer and wider than blots. Leave one end open and insert the dry blot . Wet it with 2X SSC. Pour 2X SSC out.

3. Add 5 to 10 ml Hybridization solution and seal bag. Note that bags are harder to seal when they contain the fluid, so you may have to turn up the sealing time on the heat sealer to get a good seal.

4. Incubate 68oC for 30 min. During this time, calculate how much of each probe to add to a fresh 10 mls of Ultrahybe to reach 10 to 20 ng/ml (about 0.1 nM).

5. Heat cRNA probe at 94oC for 10 min - chill on ice.

6. Cut off a corner of the bag.

7. Discard prehybe buffer. Add Ultrahybe containing the appropriate probe to each blot.

8. Reseal bag with heat sealer

9. Incubate 65oC overnight or over weekend.

10. Predict Blot results

Based on your blot photo and what you know about actin mRNA (2100 bases) and GAPDH mRNA (1400 bases), draw your expected hybridization results in your notebook. Use the 28S & 18S rRNA positions as markers. For the mouse, 28S rRNA is 4718 bases and 18S rRNA is 1847 bases. There should be 2 to 6 pg of GAPDH mRNA in 5 ug of the mouse liver RNA + control.

Q: How much RNA is in each rRNA band?

C. Assess quality of probes.

1. Make 4 serial 1:9 dilutions of each probe: Pipet l undiluted probe into a tube with 9 ul TE and mix. This is a 1:10 dilution. Repeat the dilution 3 times; so have 1:10, 1:100, 1:1000 and 1:10,000 dilutions.

2. On three 3 by 10 cm strips of nylon membrane, mark membrane with sharp pencil or black Sharpie pen "-1", "-2", "-3", "-4" at 2 cm intervals. Labelled one for each probe. Add initials and date.

3. Dot 1 ul of appropriate probe dilutions under labels. UV Cross-link RNA to membrane as you did for the Northern blot.

4. Air dry and store in a clean place. (You'll develop the dot blot along with the Northern blot in the next lab.)

Lab7 10/18

Northern Blot Washing and Development; Making cDNA by Reverse Transcription

A. Northern blot washing and development

After overnight hybridization, probe is maximally bound to specific sequences. It is also present on some non-specific sites. By reducing [salt], mainly in the form of SSC, hybridizations are tested for stringency. Usually temperature is increased as well so that probe-binding is specific for the target of interest.

KEEP BLOTS WET DURING THESE PROCEDURES or you’ll generate a lot of artifacts.

1. Cut off corner of hybridization bag. Discard hybe solution.

2. Cut bag open and move blot to a clean tupperware container. Put all 3 blots in the same container. Seal the lids during washes so solutions don’t spill onto the shakers!!!

3. Wash the 3 blots in 100 mls of 42oC 2x SSC with shaking for 15 min.

4. Discard wash and repeat.

5. Wash in 100 mls 0.5 X wash solution [Maleic acid buffer (1X = 0.1 M maleic acid, 0.15 M NaCl, pH 7.5) with 0.3% Tween-20] at 65oC for 15 min. Repeat once.

6. Use CDP-Star detection system at RT:

a. Add in the probe dot blot from Lab 6C at this step. Equilibrate membranes in washing buffer 1 min:

b. Allow chemiluminescent substrate to come to RT.

c. In freshly washed tupperware, block the membranes for 30 min in 20 ml block solution (1% (w/v) blocking reagent in maleic acid buffer without Tween). During gentle shaking, membranes should move independently from each other.

d. Discard block and incubate in Anti-DIG-Alkaline Phosphatase antibody solution (1:20,000 diluted in block solution) for 30 min. Use a minimal volume (20 ml) in a small clean container like a yellow tip box lid. During gentle shaking, membranes should move independently from each other.

e. Discard the antibody solution and wash twice in washing buffer for 15 min each time.

f. Discard wash and equilibrate membrane in detection buffer for 2 min.

g. Pipet 1 ml of CDP-star detection reagent diluted 1:100 in detection buffer onto each blot and cover with plastic wrap. Incubate 5 min, then pour off excess reagent.

7. Wrap blot in saran wrap. To keep the blot wet and the film dry, double fold the plastic wrap and tuck all edges under the blot. As always, RNA side up!

8. Place in cassette.

9. Go to the dark room and lay a piece of film on the blot. Bend the lower right corner of the film. Close the cassette. Make sure you fold the flaps so it is light tight!

10. Place cassette at 37oC or room temperature. Develop the film in 30 min. If desired, place new film on and expose longer…O/N?.

B. Making cDNA by Reverse transcription

To produce cDNA for cloning, mRNA is reverse transcribed. Typically, an oligo dT primer is annealed to the poly A tails of the mRNA's. The Reverse Transcriptase (RT), the enzyme retroviruses use to copy their RNA genomes, copies the RNA into cDNA. This reaction is used for first strand cDNA synthesis, the first step in formation of all cDNA libraries, and is the basis for true molecular cloning.

Each person will perform one RT synthesis of cDNA on their favorite RNA sample.

1. Spin down stock RNA sample after adding 1/10 vol 3M NaAc and mixing.

Dissolve in a volume of 1 mM Na citrate to make solution 1 ug/ul.

(estimate from absorbance measurements). Measure A260. Pipet 10 ug into clean tube and bring up to 25 ul volume with 1 mM Na citrate.

2. Combine in a 1.5 ml tube at RT, in order:

5.0 ul DEPC-H2O

5 ul 5X AMV RT Buffer

1 ul 100 mM DTT

2 ul dT17 primer

5 ul RNA (2 ug)

3. Heat 68oC 5 min.

4. Let cool to RT slowly.

5. Add 5.0 ul 5 mM dNTP

1 ul RNasin

1 ul AMV RT

6. Incubate 37oC, 1 h.

7. Heat at 94oC for 5 min. to kill RT.

8. Store at -20oC.

Lab 8 10/25

Analysis of Blots; Quantitative RT-PCR

A. Northern Blot Analysis

1. Develop the autoradiograph in the film processor. Label films with exposure date, time and index to your notebook. Align with the blot and mark the well, 28S & 18S rRNA positions on the film. What are the positions of the hybridized bands? Are the bands more intense in RNA samples from one tissue compared to the other?

2. Qualitatively assess the blot results:

a. Is exposure optimal?

b. How many bands are evident? Are they in the expected tissues?

c. What is the size of the darkest band? To do this measure migration distances for the band on the Xray film, and for 28S (4800 bases) and 18S (1800 bases) rRNAs on the EtBr stained gel photo using UV ruler as a guide. On graph paper, plot log base length against migration distance for the rRNAs. Find log bases from the plot, using the migration distance of the band of interest. Find antilog to get number of bases. If RNA markers show up, you can use those instead of or in addition to the 18 and 28S rRNAs.

3. Quantitate blots –

a. Can use densitometry on the Xray film. Need a good scanner and analytical software such as BioImage IQ.

b. Many machines are being developed for direct scanning of blots. These are very powerful because they avoid the limitations of film.

B. Quantitative PCR analysis of ER mRNA in an RNA sample

NOTE:Be clean (Don't introduce exogenenous DNA) USE AEROSOL BARRIER TIPS! WEAR GLOVES!

PCR amplification is very useful in detecting small amounts of nucleic acids. Since the amplification can be described mathematically in theory, then it should be possible to calculate back to the original concentration of template. Problems arise because the exponential amplification is seldom realized. To reduce variability and gain quantitative ability, PCR reactions are done with the fewest possible cycles under limiting reagent conditions. An internal competitive template, with the same primer/sites on a shorter piece of DNA, can be added in increasing amounts to a PCR reaction. In the reaction with equivalent products: large from the template of interest and small from the internal competitor; the starting amounts of template are equivalent. So by this PCR titration with competitor template and gel analysis with EtBr staining, templates of interest can be quantitated. Quantitative PCR is very well explained in Clontech's MIMIC manual.

How do you get template DNA? The quantitative PCR can be tested on linearized plasmid DNA. But to use PCR to estimate [specific mRNA] in an RNA sample, it is first reverse transcribed to cDNA as you did in the last lab.

Each person will do a set of five quantitative PCR reactions on one RT-cDNA sample, as well as (+) and (-) PCR controls.

1. At RT, combine (in order) 8X amounts of the first five reagents to make a master mix. Add 23 ul of master mix to seven 200 ul thin-walled tubes, then add the template and internal control DNAs to each individual tube.

Master mix: 17.5 ul sterile H2O X8 = 140 ul

2.5 ul 10X Taq Buffer containing Mg++ X8 =20ul

1 ul 5 mM dNTPs X8 = 8 ul

1 ul primer A (ERPCR1, 10 ng/ul) X8 = 8 ul

1 ul primer B (ERPCR2,10 ng/ul) X8 = 8 ul

2. Do serial dilutions (10E–1, 10E-2, 10E-3, 10E-4) in sterile water of the provided internal control DNA(linearized poER8short, 100 amol/ul). Add DNAs to your tubes:

TUBE #1 1 ul RT-cDNA and 1 ul 10E-4 internal control

TUBE #2 1 ul RT-cDNA and 1 ul 10E-3 internal control

TUBE #3 1 ul RT-cDNA and 1 ul 10E-2 internal control

TUBE #4 1 ul RT-cDNA and 1 ul 10E-1 internal control

TUBE #5 1 ul RT-cDNA and 1 ul undiluted internal control

TUBE #6 1 ul positive control plasmid (poER8, linearized, 1 amol/ul) and 1 ul H2O

TUBE #7 2 ul H2O [negative (no template) control]

All should have 25 ul final reaction volumes.

Note: PCR primer sequences are:

ERPCR1 - (5') agcccagcggctacaggtgc

ERPCR2 - (5') gcaggcctggcagctcttcctcct

3. Program the PCR machine to do 40 cycles of:

94oC for 30 sec: strand separation

50oC for 30 sec: primer annealing

72oC for 1 min: polymerization

4. Put your 7 PCR tubes in the PCR machine. Add 0.25 ul Taq DNA Polymerase with a P-2 Pipetteman and tips during a 94oC hot start (make machine hold at 95oC for 2 min then add enzyme. You can also do this in a hot block or bath. This "hot start" prevents lots of nonspecific product from primers annealing non-specifically with the template at RT.) Run the cycling program after enzyme addition.

5. Make one 2% agarose gel for each two students:125 mls of 2% agarose gel + 0.5 ug/ml EtBr in 1X TAE; use 2 thin combs for each gel .

6.. Store PCR reactions at -20oC. Store gel in the gel mold with the comb, wrapped in saran wrap with a little buffer and in a sealed tupperware container or pyrex dish at 4oC.

Lab 9 11/1

Quantitative RT-PCR Analysis; PCR Subcloning

A. Quantitative PCR Analysis

1. Add 2.5 ul 10x DNA dye to each 25 ul of PCR reaction. Mix by pipetting up and down.

2. Load the entire samples (or as much as possible in the well) on the 2% gel made in previous lab. Also load pGEM DNA markers (2ug = 20 ul).

3. Run at 120 V for 1 h.

4. Photograph.

5. Estimate

a. Sizes of products in bp.

b. [target cDNA] in RT-cDNA reaction.

NOTE: The ratio of the target cDNA (315 bp) to the internal control (249 bp) is the critical endpoint. Find the reaction where the two bands are nearest to being equivalent in brightness of EtBr staining, and that is the reaction that started out with equal amounts of target mRNA and internal control. Since you know how much of the latter you added, you know how much of the target was in the sample. You can now determine the concentration of mRNA in your original RNA prep.( You can assume the RT reaction made one cDNA copy of each mRNA present.)

To make this technique amenable to comparing mRNA levels in a large sample set, a single internal standard concentration equal to the average target concentration is used with each sample. Radioactive dNTP's can be used to label the products so their ratios can be quantitated. There are machines that quantitate UV fluorescence directly from gels, too.

NOTES on INTERPRETING AGAROSE GEL INFORMATION:

By comparing the bands in plasmid lanes with DNA standards, one can describe

a. The molecular size of the fragment, and

b. The [DNA] of each fragment

For (a), compare migration distance to that of DNA standard fragments - estimate bp size.

For (b), compare brightness of bands to those of DNA fragments - estimate ng. Divide by the amount of sample loaded to get [DNA]. This is often more reliable quantitation than A260 measures!!!

NOTE: MARK DYE POSITIONS ON GEL PHOTOGRAPHS. Label all photos completely! Name, date, identity!!!

B. PCR sub-cloning (see cartoon following)

One of the most useful applications of the polymerase chain reaction (PCR) is easy subcloning. Subcloning, taking a piece of cloned DNA and transferring to a new vector, used to be limited due to the scarcity of naturally occurring restriction enzyme sites. One was forced to choose DNAs with restriction sites, often these were for weird, expensive, and inefficient restriction enzymes. With PCR, restriction enzyme sites can be engineered on any DNA by adding the desired sites to the ends of the primers. Note that 5' and 3' primers must have different sites to prevent them from annealing. After restricting the PCR product, sites are different on the ends and can only enter a similarly restricted vector in one orientation: therefore, this is "Directional cloning."

Although PCR can be temperamental, subcloning fragments less than 1000 bp is, dare I say, fairly easy.

Here, we'll subclone our beta globin cDNA into a different plasmid vector. The old vector was pBluescript (Stratagene); the new pET-5a (Promega).

Q: What specific advantages does the new vector have (see Promega catalog or protocol book)? List these in your notebook.

Do 1 PCR reaction per person. Be clean (Don't introduce exogenenous DNA): USE AEROSOL BARRIER TIPS! WEAR GLOVES!

1. Combine, in order, at RT in a 200 ul thin-walled PCR tube

17.5 ul H2O

2.5 ul 10X Taq Buffer containing MgCL2

1 ul 5 mM dNTPs

1 ul primer Xba-GLOB (0.25 ug/ul)

1 ul primer GLOB-R1 (0.25 ug/ul)

1 ul linearized pGLOBcdsonly

0.5 ul Taq DNA Polymerase

Note: primer sequences are:

Xba-GLOB (5') gctctagatgctggttgtctacccatgg

GLOB-R1 (5') gcgaattctgaagttctcaggatccacg

2. Mix by pipetting up and down

3. Program a PCR machine for 35 cycles of:

94oC for 30 sec: strand separation

50oC for 30 sec: primer annealing

72oC for 1 min: polymerization

4. When the PCR is complete, the reactions will be stored at –20oC until the next lab session.

Lab 10 11/8

DNA Restriction and Ligation

A. DNA Restriction and purification on gel

Ligations are the weak point in most cloning procedures. Here we are set for success by using "sticky ends" restriction sites with 4 base overhangs that like to pair up. We'll also do the ligation directly in low melting agarose to minimize loss of DNA during purification.

NOTE: Restriction Enzyme cuts usually contain 1-2 ug DNA and 5-10 units of enzyme in 20 ul. Working buffer strength is always 1X. Use the buffer supplied with the enzyme if it is a single enzyme cut. If using two enzymes (as here), use a buffer compatible to both! Which one should you use????? There is a table in the Promega book that will help you choose.

1. Add 25 ul of TE to the PCR reaction and transfer the 50 ul to a 500 ul microfuge tube.

2. Extract with Phenol/Chloroform/IAA (25:24:1) pH 8.( This means to add an equal volume of Phenol/Chloroform/IAA, vortex to make an emulsion, microfuge 2 min., then transfer the upper aqueous phase that contains the DNA to a clean, labeled tube.)

3. Extract with Chloroform/IAA (24:1)

4. Set up a restriction digest for the PCR product:

16 ul PCR product (insert)

2 ul 10X ?????? Buffer

1ul EcoR1

1 ul Xba I

20 ul TOTAL volume

5. AT THE SAME TIME, RESTRICT 0.5 ug of the vector pET-5a (see map in Promega book) as in 4. Check calculations for the reaction with the instructor.

6. Incubate at 37oC for 1 h.

7. Make a 0.8% low melt agarose gel + EtBr. Raise comb by adding small squares of tape. These gels are like soft jello - Hard to handle!! Be careful removing combs, etc.

8. Add 2 ul 10X Loading dye to the digest reaction. Load it and 1 ug Lambda HindIII EcoR I DNA markers on the gel. Run at 100 V for 30 min.

9. Cut out bands and remove excess agarose while visualizing under Long wave UV light. Wear goggles!

B. DNA LIGATION

1. Melt DNA + agarose 70oC, 10 min. Then keep tube at 37oC until through pipetting.

2. Label two ligation tubes. Put them at 37oC. Add 1 ul vector DNA in gel to each. To the ligation tube, add 3 ul insert DNA in gel and 5 ul H2O. To the control tube add 8 ul H2O. Keep these at 37oC for at least 2-3 min and until the master mix is added.

3. Make a Master mix composed of:

2 ul 10X Ligase Buffer x 3 = 6 ul

8 ul H2O x 3 = 24 ul

1 ul T4DNA Ligase x3 = 3 ul

11 ul per reaction

Add 11 ul ice-cold mix to each ligation tube. Finger flick the tube immediately and slam on ice. Reactions will gel while ligation occurs.

4. Incubate 15oC, O/N. This is done in a hot block in a 4oC room.

Lab 11 11/15

Transformation of Bacteria

"Bacterial transformation" relates to the change of bacterial phenotype by introducing a plasmid containing an antibiotic resistance gene. "Competent cells" are made receptive to plasmids by making their membranes permeable with calcium treatment. Plasmids adhere to cells, enter on heat shock, and cells are selected for antibiotic resistance on plates after a 1 h recovery period in broth. NOTE: All waste contaminated with E. coli must be killed with bleach or discarded in "BIOHAZARD BAGS," which are autoclaved prior to disposal.

Each group does three transformations: one from the vector only control ligation and one from the ligation that has DNA insert too, as well as a positive control for transformation: 10 ng of circular plasmid. For the last, use any plasmid.

1. Thaw cells on ice (20 min.)

2. Pipet 100 ul into a cool 1.5 ml tubes: 1 for each ligation (vector only and vector+insert), 1 for the positive control

3. Melt ligations at 70oC, 10 min.

4. Cool ligations to 37oC in block, at least 2 min.

5. Add 1 ul ligation to cells; mix by pipetting up and down.

6. Incubate on ice 30 min. Mix every 10 min. by tapping th tube gently.

7. Heat shock 42oC, 45 sec (Be exact here!). No shaking.

8. Ice for 2 min.

9. Add 400 ul room temperature S.O.C. broth.

10. Agitate cultures gently, 37oC, 1 h. Tube turners or rockers are good for this.

11. Spread all 500 ul on a LB + Ampicillin plate for the transformations from ligations. Spread only 50 ul for the transformation of circular plasmid.

12. Incubate 37oC O/N. Invert plates if all the liquid goes into the plate. Incubate wet plates right side up.

Thursday 11/21 (any time between 2 to 3 p.m.?)

1. Count colonies on the ligation plates. Compare to the number of colonies on vector only control plate. If you have twice as many colonies in the ligation transformation plate, then 50% should contain the insert!

2. Make 4 - 10 ml overnight cultures of LB + Amp in 50 ml tubes. Label tubes 1-4 (initials and date).

3. Inoculate @ with 1 colony: from vector only plate (1 culture) or from ligation plate (3 cultures).

4. Incubate with strong agitation (200 rpm) at 37oC O/N.

Lab 12 11/22

Plasmid DNA Miniprep

NOTE 1: Phenol for DNA is buffered with Tris to pH8 for optimal partitioning. For RNA, Phenol is water-saturated and is ~pH5. Use the correct phenol for your nucleic acid!!!

NOTE 2: ALSO, Phenol is corrosive - causes burns! Be careful! Wear safety glasses!!!

NOTE 3: CHCl3 (Chloroform) dissolves things like styrofoam and polystyrene - use glass graduated pipets and polypropylene 15 & 50 ml tubes.

In a short, mind-numbing period of tube shuffling, one can extract plasmid DNA for analysis of desirable clones

1. Spin down 1.5 ml overnight culture cells in a 1.5 ml tube - (DISCARD all materials from steps 1 -3 that are contaminated with cells in BIOHAZARD BAGS.)

2. Save the rest of the overnight culture at 4oC - IF it is a good clone, you'll want to make a glycerol stock for long term storage and a streak plate for short term use.

3. Remove the medium by aspiration, leaving the bacterial pellet as dry as possible.

4. Resuspend the pellet by trituration (pipetting up and down) in 150 ul of an ice-cold solution of:

50 mM glucose

10 mM EDTA

25 mM Tris/ HCl (pH 8.0)

Add 4 mg/ml lysozyme (added freshly to the solution)

5. Store for 5 minutes at room temperature. The top of the tube need not be closed during this period.

6. Add 300 ul of a room temperature solution of: 0.2 N NaOH + 1% SDS

Close the top of the tube and mix the contents by inverting the tube rapidly two or three times. Do not vortex. Store the tube on ice for 5 minutes.

7. Add 225 ul of an ice-cold solution of potassium acetate (~pH 4.8). [This reagent was made up as follows: To 60 ml of 5 M potassium acetate, add 11.5 ml of glacial acetic acid and 28.5 ml of H2O. The resulting solution is 3 M with respect to potassium and 5 M with respect to acetate.]

Close the cap of the tube and vortex hard, put on ice, vortex again. Store on ice for 5 minutes.

8. Centrifuge for 15 minutes in an micro-centrifuge at 20oC.

9. Transfer 600 ul supernatant to a fresh tube. (Avoid all white, solid garbage).

10. Add an equal volume of phenol/chloroform pH8. Mix by vortexing. After centrifuging for 2 minutes in an Eppendorf centrifuge, transfer the aqueous phase to a fresh tube.

11. Add two volumes of ethanol at room temperature. Mix by vortexing. Stand at room temperature for 2 minutes.

12. Centrifuge for 5 minutes in an Eppendorf centrifuge at room temperature.

13. Remove the supernatant. Stand the tube in an inverted position on a paper towel to allow all of the fluid to drain away.

14. Add 1 ml of 70% ethanol. Vortex briefly and then centrifuge.

15. Again remove all of the supernatant. Air dry the pellet briefly (5 minutes) after wiping away residual ethanol with a Kimwipe.

16. Add 50 ul of TE (pH 8.0). Vortex and incubate at 37oC to solubilize DNA (about 5 minutes).

17. Store at -20oC until next lab session.

Lab 13 12/6

Miniprep plasmid DNA Restriction and Gel Analysis

A. To analyze the four plasmid preps from overnight cultures, we'll restrict each with EcoRI and Bam HI to see if the 400bp insert is present.

1. We will cut inserts out of the vectors for each plasmid prep to verify it. Best pipetting technique minimizes pipetting steps (and, thus, work and error). So, to set up a set of 20 ul restriction digestions, make a master mix of common components:

In single reaction x5 = in master mix

6ul H2O x5 =30 ul

2 ul 10X????? buffer x5 =10ul

1ul 1mg/ml RNase x5 =5 ul 1mg/ml RNase

0.5 ul Bam H1 (10 U/ul) x5 = 2.5 ul Bam H1

0.5 ul EcoR1 (10U/ul) x5 = 2.5 ul EcoR1

Use 10ul/rxn

Mix gently. Pipet 10ul into 4 tubes.

Add 10 ul of individual plasmid preps, one to each tube. Incubate 1 h at 37oC. ALSO Prepare one uncut plasmid sample: 10 ul plasmid + 10 ul TE and incubate at 37oC. After incubations, add 2 ul 10x DNA dye to each of the five tubes. Store leftover plasmid at -20oC.

2. Make a 1% agarose gel + 0.5 ug/ml EtBr in 1X TAE. Use a mid-size apparatus and two 14-well combs.

3. Add 2 ul 10X DNA dye to @ digest.

4. Load gel with uncut plasmid (1 sample) and 4 plasmid digests. Also load Lambda Hind III EcoR1 markers (1 ug) in one well of top and bottom halves of each gel.

5. Run at 120V, 1 h.

6. Photograph under UV light.

7. By comparing the bands in plasmid lanes with Lambda standards, one can describe

a. The molecular size of the fragment

b. The [DNA] of each fragment, and

c. the molecular form (circular vs. linear) of the fragments

For (a), compare migration distance to that of Lambda standard fragments - estimate bp size.

For (b), compare brightness of bands to those of Lambda fragments - estimate ng. Divide by 10 ul (amount of sample loaded) to get [DNA]. This is often more reliable quantitation than A260 measures!!!

For (c), note that uncut plasmids run fast as two forms, supercoiled and nicked circular. The restricted plasmid from vector only transformation shows that linear DNA is slower. Since markers are linear DNAs, their migration only relates to other linear DNAs.

NOTE: MARK DYE POSITIONS ON GEL PHOTOGRAPHS. Label all photos completely! Name, date, identity!!!

Lab Clean-up:

Biohazard waste

Discard phenol in appropriate bottles

Samples - save?

Return: 10 ml pipettors

APPENDIX

Use of Micropipettors

1. Choose the correct pipet. For volumes:

1-20 ul P20

20-200 ul P200

200 - 1000 ul P1000

2. Set the desired volume by holding the pipetman in one hand and turning the volume adjustment knob until the correct volume shows on the indicator. For best precision, always approach the desired volume by dialing downward (at least one-third revolution) from a larger volume setting.

3. Attach a new tip to the shaft of the pipet. Press tip on firmly to ensure airtight seal. Chose the correct tip.

P20 yellow tip

P200 yellow tip

P1000 blue tip

4. Depress plunger to first positive stop. Hold pipetman vertically and immerse disposable tip into sample liquid 2mm.

5. Allow the push button to return slowly to the up position. Never permit it to snap up.

6. Wait 1 or 2 seconds to ensure that the full volume of the sample is drawn into the tip.

7. Withdraw tip from the sample liquid. Wipe the sides of the tip on the sides of tube to remove any remaining liquid.

8. To dispense the sample, place the tip end against the side wall of the receiving vessel and depress the plunger slowly to the first stop. Then depress the plunger to the second stop to expel any residual liquid in the tip.

9. With the plunger fully depressed, withdraw pipetman from the vessel. Then allow the plunger to return to the top position.

10. Discard tip by depressing the tip ejector button. A fresh tip should be used for each sample.

Terribly Difficult Calculations

1. Molar solutions

1 M (mole per liter) means the solution has 1 molecular weight mass (g) per volume (liter) of soln.

A mole is a number of molecules:

6.022 x 10E23, Avogadro's number

To make 500 mls of 0.5 M NaCl (NaCl is58.55 g/mole) you need (0.5 liters)

(0.5 mole )= 0.25 mole

liter )

0.25 mole * 58.55 g/mole = 14.6 g

So: Add 14.6 g NaCl power and bring final volume to 500 ml with H2O.

2. We typically work with concentrated stock solutions. For example, our Tris/acetate/EDTA (TAE) is made as a 50X stock. We run gels in 1000 mls of 1X TAE. The way I do DILUTION PROBLEMS is:

[Stock] * y = [Desired] * Desired volume; where y is the volume of stock. To find Y needed to make 1 li of 1X TAE from a 50X stock:

50X * y = 1X * 1000 ml

y = 1X/50X * 1000 ml = 20 ml

So add 20 ml 50X TAE to a 1 liter graduated cylinder. Bring volume to 1 li w/ dH2O.

3. Note: Dilutions are applicable to problems of pipetting very small amounts. If you want to add 0.2 ul, dilute the material 1:9 and pipet 2 ul with a P-20.

4. Percentage solutions should have a (v/v) or (w/v) or (w/w) following.

a. (v/v) relates volume to volume, indicating both components are liquids: e.g. 100 mls of 75% (v/v) EtOH is made with 75 mls EtOH + 25 ml H2O

b. (w/v) indicates solid to liquid ratio: e.g., 10 mls of 10% (w/v) ammonium persulfate (APS) is made w/ 1 g of APS to 10 ml final volume with water.

c. (w/w) is rare, indicating a weight to weight relationship. To make 10 mls of a 10% (w/w) APS soln, you could weigh 1 g APS on a scale and then add water until solution weight is 10 g. (That would be 9 g = 9 mls since density of H2O is 1 g/ml).

5. Of course, these calculations can be combined. For example, to make 500 mls of 0.5 M NaCl in 1X TAE,

Combine 14.6 g NaCl with 10 mls 50X TAE. Bring volume to 500 mls with H2O.

Easy!

PROBE TEST GEL

(short, fat sequencing gel)

5% acrylamide/urea gel

25.5g urea

19.5ml H2O

12ml 5X TBE

7.5ml 40% (w/v) acrylamide (19:1 acryl:bis)

60ml final vol.

Heat to 37oC to dissolve urea

Cool to below RT

Filter (~optional)

Add 400 ul 10% APS (less than 1 week old, make 1ml)

50 ul TEMED