Blackboard Notes Day 1 (Mon)

READ ALL OF THIS SHEET BEFORE PROCEEDING

You will begin the course by carrying out a Western blot to determine the molecular weight of b-actin. You will start by preparing a protein extract of HeLa cells and determining the protein concentration using a dye binding assay.

Next these proteins will be separated by SDS PAGE (Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis) along with protein molecular weight standards and transferred by electrophoresis to a PVDF (polyvinylidene difluoride) membrane (Immobilon). The membrane will be stained to demonstrate the pattern of total cell protein and to ensure that all areas of the gel transferred properly.

Next, you will continue the Western blotting to identify the protein b-actin on the blots by using a monoclonal antibody specific for b-actin. The primary antibody will be allowed to bind overnight . Tomorrow, the bound primary antibody will be detected using a secondary antibody, which specifically binds the primary antibody. The secondary antibody is linked to the reporter enzyme horseradish peroxidase. Where the primary/secondary antibody/HRP complex has bound will be determined by the incubation of the membrane with a chemiluminescent substrate. Comparison of the position of the HRP reporter band relative to the molecular weight standards will allow you to determine the molecular weight of b-actin.

As time allows there will be a lecture on basic microbial techniques commonly used in molecular biology followed by a demonstration. Next there will be a lectures on phage lambda and on cDNA libraries.

Finally there will be a brief lecture on antibody screening of phage expression libraries. The procedures used in antibody screening are very similar to those employed in Western Blotting. The antibody screening process will be carried out as a demonstration in parallel with the Western Blotting procedure.

Western Blotting

Preparing Protein Extract For Western Blot

1) Use Protocol 1.1 to prepare a protein extract using the following supplemental information.

You will spin down 15 ml of HeLa cells (grown to a density of about 5 x 10⁵ cells/ml) in a 15 ml polypropylene centrifuge tube at ~750 - 1000 x g (2000 rpm IEC centrifuge) for 10 minutes in the table top centrifuge as noted in step 2 of the protocol. Pour off the bulk of the supernatant. The cell pellet and residual media will have a volume of 100-200 ml. Vortex to resuspend the pellet completely.

2) Transfer 100 ml of the resuspended cells to a screw top 1.5 ml microfuge tube, and add 100 ml 2X sample buffer at room temperature, mix and immediately float the tube in a boiling water bath for 5 minutes. Sonnicate the samples for a few seconds on medium setting to reduce viscosity of the lysates and make pipetting easier.

Determining The Protein Concentration Of The Protein Extract

1) Use Protocol 1.2 to determine the protein concentration of your extract. Add 5 ml of your sample to 95 ml of water and add this 100 ml diluted sample to 5 ml of the diluted BioRad reagent. If the color of the sample is too dark (off the standard scale) dilute a 5 ml aliquot of your sample with 15 ml of 1 X sample buffer and add 5 mof l this to 95 ml water and repeat the reading.

The standard curve has been done for you, and we will distribute the results to each student so you can estimate the protein concentration in your sample.

Running An SDS-PAGE Gel

1) Next separate proteins by size on an SDS PAGE gel using Protocol 1.3.

Note: The protocol provides a good deal of information specific to the Hoefer gel electrophoresis unit, much of which, though useful for understanding how to pour various polyacrylamide gels, is not absolutely necessary at this point because you will be provided with a pre-poured Novex or Promega 12% SDS-PAGE gel which has a 4% stacking gel on top. You will also be given multi-colored (Rainbow markers- Amersham) and unstained protein molecular weight markers (Gibco/BRL

Note: The rainbow protein molecular weight markers come in a buffer containing 50% glycerol. You will need to add an equal volume of 2x sample buffer in order for the markers to migrate similarly to the protein extract. Do NOT boil the rainbow markers as this will lead to the dissociation of the dye. Note also that the rainbow markers are not stable in the presence of b-mercaptoethanol which is present in the sample buffer and will disassociate the dye from the proteins. Therefore, do not let the rainbow markers sit in sample buffer for more than an hour before loading on a gel.

You should prepare samples for loading on the gel according to the following scheme.

Lane 1: 10 ml of the Rainbow protein molecular weight standards + 10 ml 2x sample buffer

Lane 2: 20 ml XX we can probably go back to 10 ul which will be more accurate i.e. all the same volumes, as the markers were plenty dark. )unstained low molecular weight protein markers (Gibco/BRL. Note These standards come in a storage buffer similar to our sample buffer. Hence there is no need to add sample buffer to these standards. Note also you are adding four times the recommended amount for gel detection of the markers, but since the transfer of the proteins and their binding to the Immobilon P membrane may not be quantitative, this higher amount should ensure adequate staining after transfer.)

Lane 3: 30 mg of your protein extract (Student 1)

Lane 4: 10 mg of your protein extract (Student 1)

Lane 5: 3 mg of your protein extract (Student 1)

Lane 6: 10 ml of the Rainbow protein molecular weight standards + 10 ml 2x sample buffer

Lane 7: 20 ml of the unstained low molecular weight markers (see note in step 2)

Lane 8: 30 mg of your protein extract (Student 2)

Lane 9: 10 mg of your protein extract (Student 2)

Lane 10: 3 mg of your protein extract (Student 2)

Note: You should be able to calculate the volume of cell extract you will need for each lane from the protein concentration you determined earlier. Your sample volume per well should be kept to approximately 20 ml or less, otherwise the sample will exceed the capacity of the well and spill over into the next lane. The samples will be easier to load, and spill over will be less of a problem if the final sample is ~10 ml. Another consideration is that the gel will run more evenly if the final volume of all samples, including the markers, are the same. Thus, you should first determine how many mls of your sample correspond to 30 mg of protein. If your sample is so dilute that 30 mg is in a volume of greater than 18 ml, then load what you can in 18 ml and record the amount. If the sample is very concentrated, then dilute to 9 ml with 1X Sample Buffer, made by diluting 2X Sample Buffer with water. Once this volume of your highest concentration sample has been established, bring all the other samples to the same volume with 1X sample buffer.

2) Because our sample loading buffer lacked the electrophoresis tracking dye bromophenol blue which can interfere with the protein dye binding assay, you will now have to add 1/10 volume of 1% bromophenol blue to each of your extract samples. Tap to mix.

3) Now mark the bottom of each well with a marker and CAREFULLY remove the comb from your prepoured Promega or NOVEX gels. The TAs will demonstrate how to assemble the gel apparatus using Promega or NOVEX gels and the Hoefer SE250 electrophoresis rigs. Proceed with steps 4 and 5 of the protocol. Then skip steps 6 to 8 and proceed to step 9. The proper way of underlaying a sample in a well will be demonstrated.

4) Your group and the other group in your quadrant will share a single gel apparatus. Run your gels at 125 V constant voltage. The initial current will be about 40 mA for the two gels (20 mA per gel). Run the gels for about 90 min. until the bromophenol blue marker is at the bottom of the gel. Be sure to observe the correct polarity of your electrophoresis apparatus i.e. the anode (red) the positive terminal at the bottom. (Question: Why do all the proteins move toward the anode?) Note how the samples stack as they enter the stacking gel, and how the colored molecular weight markers separate during the electrophoresis.

5) At the completion of the electrophoresis turn off the power and disconnect the gel box. Open the Novex cassette by breaking the tabs with a gel knife. The gel should remain on the un-notched plate (labeled "12% TG"). Next use the gel knife to cut away the gel tab at the base of the cassette. Finally cut off the vertical protruding wells at the top of the gel before removing the gel from the cassette. XX You may also wish to cut off the stacking gel at this point. In homemade gels the stacking gel will often spontaneously separate from the resolving gel or at least the interface between the two gels will be easy to see. However in these commercial gels you will probably be unable to see the boundary between the two gel types. Novagen informs us however that the height of the stacking gel is about 8 mm, so you can measure off this distance from the bottom of the wells and make your cut there. Although we should have no trouble orienting the gel because of the asymmetrical loading pattern, you might want to cut off the upper left corner edge of the gel so that you can immediately orient the gel.

6) Using a wash bottle or the spatula, free the gel from the plate and allow it to fall into a dish of water. You may find that water from a squirt bottle can help to dislodge the gel. Then carefully pour off the water and add buffer II (Protocol 1.4) so that the gel will have a chance to equilibrate with the buffer prior to electrophoretic transfer of the proteins to the membrane.

Transfer Proteins From SDS PAGE Gel To A Membrane

1) You next need to transfer your separated proteins to a PVDF (Immobilon P) membrane filter exactly as described in Protocol 1.4. Be sure to prewet the Immobilon filter in methanol and then water as noted in the protocol. The assembly of the gel/membrane sandwich will be demonstrated by the TAs. You should note the orientation of your gels on the membrane e.g. lane 1 to the left so that when you finally develop your film after antibody screening you will be able to readily align your film to the blot.

Note: Each group of two students who shared a single gel will assemble a single transfer sandwich as shown in the figure. The two gels from each quadrant will be placed side by side in the electroblotter to form a single layer. After the first quadrant of students has assembled their two gel sandwiches, place a wet sheet of dialysis membrane on top of the sandwiches. The second quadrant of students will then construct a second layer of sandwiches on top of the first. The dialysis membrane will serve to ensure that no proteins from the upper layer migrate through to the lower layer while permitting the free flow of electrons.

Note: In step 5 you must use the total area of both gels in a single layer in determining the total current to use. (You need not consider the area of the gels in the second layer of sandwich since the gels are "in series").

2) After the completion of transfer, disassemble the blotting apparatus. You should see the colored protein markers nicely transferred to your filter. The identity and molecular weight of these markers is as follows:

Myosin (M.W. 220,000) Blue

Phosphorylase B (M.W. 97,400) Brown

Bovine Serum Albumin (M.W. 66,000) Red

Ovalbumin (M.W. 46,000) Yellow

Carbonic Anhydrase (M.W. 30,000) Orange

Trypsin Inhibitor (M.W. 21,500) Green

Lysozyme (M.W. 14,300) Magenta

Note: The transfer of the Rainbow markers to the blot can indicate the quality of your transfer. Hopefully you will see that bands of all molecular weight transferred roughly equally. Cut off the upper left corner of the filter as an aid to subsequent alignment. At the edge of the membrane mark the location of the markers with a pencil in case they fade during subsequent incubations. Note the color of the markers at each location at this time, because after staining in the next step, all the Rainbow markers will stain blue. Dry the filter using the methanol procedure as given in the protocol to fix the proteins to the membrane.

Rewet the membrane in methanol followed by water, then stain and photograph the membrane for total protein to determine transfer efficiency following Protocol 1.5. It is very important that you reequilibrate the dried Immobilon membrane in methanol and then water prior to carrying out the staining. The filter is highly hydrophobic and will not wet in aqueous solutions without prior equilibration in methanol. (Note that nitrocellulose filters which are occasionally used for Western blotting are naturally hydrophilic and do not require wetting in methanol, in fact they will dissolve in methanol.)

After staining you should be able to see the previously unstained Gibco/BRL molecular weight protein markers following the 20% methanol destaining step. Mark their position on the blot with pencil as well. The unstained Gibco/BRL markers are as follows:

Unstained Gibco/BRL markers

M.W. 220 M.W. 40

M.W. 160 M.W. 30

M.W. 120 M.W. 25

M.W. 100 M.W. 20 Highlighted (twice as dark as surrounding bands)

M.W. 90 M.W. 15

M.W. 80 M.W. 10

M.W. 70

M.W. 60

M.W. 50 Highlighted (twice as dark as surrounding bands)

Note: Some of the lowest M.W. markers may have run off the end of the gel.

a) Check your stained filters. Did the transfer appear to be uniform?

b) After photographing the filter, be sure to destain as noted in the Protocol 1.5, but note that destaining will probably not be complete. This is of no consequence to subsequent detection by ECL.

Immunodetection Of b-Actin Protein

1) Proceed with immunodetection Protocol 1.6 up to and including step 5. Do the blocking step (~30 minutes) in 50 ml PBST 5% nonfat dry milk and the washes in cold PBST. Let the membrane incubate overnight at 4^oC in 25 ml primary antibody which is a monoclonal anti actin (Amersham) diluted 1:5000 in PBST 5% nonfat dry milk. You may wish to refer to the Amersham manual that contains excellent diagrams of how the chemiluminescent detection procedure works. Incubations and washes are conveniently carried out in 150 mm Petri dishes.

Demonstration Of Sterile Technique and Dilution Procedures

1) As time allows, we will demonstrate how to use pipettes properly, how to remove solutions from a bottle without introducing contaminants, how to prepare dilution series of bacteria or phage, how to streak a bacterial culture to obtain single colonies, how to spread a bacterial culture on a plate so that individual colonies can be enumerated, and how to assay a phage stock by using the top agarose plate method. These techniques are described in Chapter 2 of the manual.

Lecture On phage lambda

Lecture On cDNA Libraries

Lecture On Screening Of Antibody Expression Libraries And A Brief Demonstration Of Titering Phage.

1) How to titer a phage library will be demonstrated.

2) You will start a 5ml O/N culture of XL1 Blue MRF' cells for use in your own titering experiment tomorrow.

Blackboard Notes Day 2 (Tue)

READ ALL OF THIS SHEET BEFORE PROCEEDING

Today you will begin by developing your Western blot to determine the molecular weight of b-actin. During the washing steps, Wayne will be completing the antibody screen of a cDNA library in parallel. Upon completion of the Western blotting procedure there will be a discussion of both the Western Blotting and Antibody Library screening procedures. Next there will be a lecture on phagemid rescue. Subsequently you will plate some lambda phage containing the b-actin insert following the procedures that were demonstrated yesterday. These are the same phage you will use in the phagemid rescue experiment that you will start next. Because the plates you are plating will not be ready until tomorrow, you will be provided with plates already containing plaques in order to begin the phagemid rescue experiment today. Finally, there will be a lecture and demonstration of "color selection" using the lacZ system.

Western Blotting (Continued)

The first thing you should do today is to pour off your diluted primary Ab.

Note: If you were doing this experiment in your lab, it would be possible to save this diluted Ab solution by mixing it with an antibacterial agent such as 0.1% sodium azide and storing the solution at 4^oC.

2) Wash your blots as described in Protocol 1.6 step 6, and incubate your blots for 1 hour at room temperature in the secondary antibody diluted 1:10,000.

3) While your blot is incubating with the secondary antibody diluted we will talk about enhanced chemiluminescence. You may also wish to look at the diagrams of the chemiluminescence procedure in the Amersham or Pierce literature to gain a better understanding of how enhanced chemiluminescence works.

4) Following the lecture, wash your blots to remove the secondary antibody and carry out ECL detection using the Pierce reagents. You may want to look at Protocol 1.6 for guidance on the detection. These procedures will be demonstrated to you.

Note: We will use sheet protectors instead of Saran Wrap for the detection steps and will soak the filters in Pierce SuperSignal detection reagent using the batch method as described in the protocol.

5) When your number is called proceed to the darkroom in groups of four and a TA will help you to put the light emitting reagents on your membrane, then expose and develop an autoradiograph. To do this you must first mix the Pierce Super-Signal Reagents A and B in equal amounts in a clean lid from a Petri plate or pipette tip box. Be sure to use a clean pipette each time you remove either one of these reagents. Cross contamination will cause these reagents to rapidly degrade. Place the blot in the mixed Pierce substrate solution for approximately 1 min.

11) After lunch we will discuss your Western blotting results.

Do the Rainbow markers and the unstained markers fall along the same line? From the position of the b-actin band on this calibration curve you should be able to determine the molecular weight of b-actin.

Screening Of cDNA Libraries Demonstration (Continued)

1) The antibody screening of cDNA libraries (Protocol 3.1) will be continued in parallel with the Western Blotting procedure to highlight the similarities between the two procedures.

2) After the Western blotting discussion we will do an exercise on how to identify lambda plaques expressing the protein of interest.

Practice Titering Lambda Phage

1) The TAs will provide overnight culture of E. coli XL1 Blue MRF'D(mcrA)183 D(mcrCB-hsdSMR-mrr)173 endA1 supE44 thi-1 recA1 gyrA96 relA1 lac [F' proAB lacI^qZDM15 Tn10 (Tet^r)]) grown in LB containing 15 mg/ml test for plating (Protocol 3.2). The cells were prepared by pelleting at 4^oC for 10 min. at 2500 rpm in the IEC centrifuge. The supernatant was poured off and the cells resuspend in 2.5 ml (1/2 original volume) of a cold 10 mM Mg⁺⁺ solution made from either MgSO₄ or MgCl₂.

Note: The cell density at this step is not terribly critical, though for optimal results the OD₅₅₀ should optimally be between 0.45 to 0.55. In reality, as long as you have a reasonably fresh culture of plating bacteria, you should be able to see the lambda plaques following plating. What is important is that the plates used for plating be reasonably fresh. Very old dry plates will yield tiny plaques that may be hard to see. If the plates are too fresh i.e. less than a day old, they may be so wet that the plaques will smear. When the plates are too wet, they can be dried in a sterile hood or incubator for a few hours with the lids slightly ajar.

2) Next titer the provided phage stock using Protocol 2.12. (See Section 2.3 for useful information on dilution methods).

3) You will be provided with 15 ml of l phage carrying the b-actin insert. Make the following sterile dilutions:

10 ml of phage stock to 90 ml SM Buffer (Dilution A)

10 ml of phage stock A to 90 ml SM Buffer (Dilution B)

10 ml of phage stock B to 90 ml SM Buffer (Dilution C)

10 ml of phage stock C to 90 ml SM Buffer (Dilution D)

You will infect 100 ml of XL1 Blue MRF' plating cells with 10 ml of dilutions A-D.

Tomorrow you will calculate the titre of this stock.

Lecture On Phagmid Rescue

Phagemid Rescue

1) Next, the teaching assistants will provide you with a plate on which they have plated a pure culture of Lambda Zap II phage containing a b-actin insert at a low enough density so that it will be easy to pick an isolated plaque. See the introduction to Blackboard Notes Day 3 to understand why we are doing this. Protocol 3.5 has a diagram explaining how the phagemid rescue works. Note that this b-actin insert has been cloned into Lambda Zap II out of frame so that no actin is expressed from this insert even if the b-galactosidase promoter is slightly leaky-which it is. Note that while we can not detect this insert using antibody techniques because no b-actin is being expressed, it is easily detectable using actin nucleic acid probes.

2) Core one of the plaques with a Pasteur pipette or wooden probe from the lawn infected with Lambda Zap II containing a b-actin insert. Elute phage from the plug in 0.5 ml of SM. In the case of the wooden probe swirl the probe in 0.5 ml of SM for several seconds and discard the wooden probe. Store the SM solution at 4^oC overnight. Tomorrow we will begin the “rescue” of the phagemid carrying the b-actin insert from this Lambda Zap II clone (Protocol 3.5).

3) The TAs will start a 5 ml overnight culture of the E. coli XL1 Blue MRF’ strain in LB + 15 mg/ml tetracycline. At the same time they will start a large culture of the E. coli SOLR strain in LB + 50 mg /ml kanamycin. Both strains are needed for the rescue on day three. (The reasons for using these particular strains for the rescue will be discussed. You should look at the following genotypes to see why these antibiotics are used).

Very IMPORTANT Note: You should always plate the XL1 Blue MRF' and SOLR cells on selective media plates and pick a single colony for this work. This 1) helps ensure that the cells have retained any necessary selective marker, particularly in the case of an antibiotic resistance gene that is carried on a plasmid. 2) It helps insure you actually have the correct cell that is not contaminated by another cell type that could interfere with the results. 3) Good colony growth is often correlated with healthy cells that will grow and perform better in subsequent liquid cultures.

XL1-Blue MRF'strain: D(mcrA)183 D(mcrCB-hsdSMR-mrr)173 endA1 supE44 thi-1 recA1 gyrA96 relA1 lac [F' proAB lacI^qZDM15 Tn10 (Tet^r)]

Note: The XL1-Blue strain allows blue/white color screening and single-strand rescue of phagemid DNA. The EndA– phenotype allows preparation of high-quality plasmid DNA. The F' episome carries antibiotic resistance, eliminating the need to select on minimal media plates. To allow high efficiency and representational cloning of methylated DNA, Stratagene created the restriction-minus XL1-Blue MRF' competent cells.

SOLR: e14-(McrA-) D(mcrCB-hsdSMR-mrr)171 sbcC recB recJ uvrC umuC::Tn5 (Kan^r) lac gyrA96 relA1 thi-1 endA1 l^R [F’ proAB lacIqZDM15]^c Su^- (nonsuppressing)

^C is an uncharacterized mutation that enhances a-complementation to give a more intense blue color on plates containing X-gal and IPTG.

The ExAssist Interference-Resistant Helper phage is an M13 MP8 derivative with none of the intragenic region being changed. There are amber mutations in genes 1 and 2.

Map Of pBluescript II KS Plasmid

T3 primer

5' AATTAACCCTCACTAAAGGG 3'

EcoO109I HincII

DraII AccI Bsp106I EcoRV EcoRI

MET BssHII T3 promoter+1 KpnI ApaI XhoI SalI ClaI HindIII

5' GGAAACAGCTATGACCATGATTACGCCAAGCGCGCAATTAACCCTCACTAAAGGGAACAAAAGCTGGGTACCGGGCCCCCCCTCGAGGTCGACGGTATCGATAAGCTTGATATCG 3'

CCTTTGTCGATACTGGTACTAATGCGGTTCGCGCGTTAATTGGGAGTGATTTCCCTTGTTTTCGACCCATGCGCCGGGGGGGAGCTCCAGCTGCCATAGCTATTCGAACTATAGCTTAA

| | |

816 792 759

b-Galactosidase→

SmaI SpeI EagI BstXI

PstI BamHI XbaI Not I SacII SacI BssHII

AATTCCTGCAGCCCGGGGGATCCACTAGTTCTAGAGCGGCCGCCACCGCGGTGGAGCTCCAATTCGCCCTATAGTGAGTCGTATTACGCGCGCTCACTGGCCGTCGTTTTACAA 3' (+)

GGACGTCGGGCCCCCTAGGTGATCAAGATCTCGCCGGCGGTGGCGCCACCTCGAGGTTAAGCGGGATATCACTCAGCATAATGCGCGCGAGTGACCGGCAGCAAAATGTT 5' (-)

l ←+ 1 T7 promoter l

657 619

3' CGGGATATCACTCAGCATAATG 5'

T7 Primer

Brief Lecture On Identification of Recombinant Vectors Carrying The LacZ System

1) Demonstration of color selection using LacZ system and estimation of the number of recombinants in a library.

Blackboard Notes Day 3 (Wed)

READ ALL OF THIS SHEET BEFORE PROCEEDING

First you should calculate the titer of your b-actin-containing phage stocks you plated yesterday. Next you will perform a basic PCR reaction using four different DNA templates. Next we will demonstrate how to pour, load and run an agarose minigel. After the PCR reaction is complete, the PCR products will be separated by electrophoresis on an agarose gel and identified using the ethidium bromide staining method. Next we will continue the excision of the b-actin-containing phagemid which is a continuation of the cDNA library antibody screening procedure which Wayne demonstrated yesterday..

cDNA Screening/Phagemid Rescue (Continued)

1) Calculate the phage titer of the cDNA phage stocks you assayed yesterday.

PCR Amplification of the Human b-actin Gene

Introduction:

In the following experiment each student will amplify a portion of the b-actin gene using b-actin specific primers. The positions of the b-actin primers we will be using are indicated on the genomic b-actin sequence (Fig. 1) and the cDNA b-actin sequence. Fig. 2. Is the size of the amplified product the same whether you use genomic or cDNA? Why or why not?

Experiment

1) You will be carrying out PCR with the following four templates:

A) b-actin pBluescript plasmid1(10 pg in 30 ml water) purified using Protocol 4.5.

B) Genomic HeLa DNA template (200 ng in 30 ml water) purified using Protocol 4.4.

Each person will be provided with templates A and B. You will have a chance to use Protocols 4.4 and 4.5 yourself later in the workshop.

C) Genomic DNA isolated from your own cheek cells using (Protocol 4.26)

D) Genomic DNA isolated from HeLa cells using the GeneReleaser microwave technique (Protocol 4.27).

Each person should carry out protocols C and D. In Protocol C you will use 30 ml of the final cheek cell extract in the PCR reaction. In Protocol D, you will take 20 ml of the final extract and add 10 ml of sterile deionized water.

Thus you will have 30 ml of each of the four templates. When you are finished preparing the four templates, you should prepare a master mix for 5.5 reactions. This amount of Master Mix will allow you to carry out 4 PCR amplification using the templates you have prepared, one no DNA control reaction, with half a reaction left over to allow for variability in pipetting.

2) Prepare a master mix for PCR enough for 5.5 reactions each of 100 ml. The polymerase we will be using is the Sigma Jump Start version of Taq which allows one to carry out an easy hot start procedure. It's a good idea in assembling the reaction to add the components in roughly the order shown. If for example you added the dNTPs directly to the 10 x PCR buffer you might precipitate the dNTPs in the high Mg of the 10 X buffer (though they probably would go back into solution later). Also, it would not be a good idea to add the enzyme to pure water in the absence of buffer. Finally note that in any PCR experiment it’s a good idea to add the template last to minimize the opening of tubes containing template that might lead to contamination of other tubes with foreign template DNA.

Per 100 ml Reaction Per 5.5. Reactions

53 ml sterile water 291.5 ml

10 ml 10X PCR Buffer 55

2 ml b actin forward primer (10 mM) 11

2 ml b actin reverse primer (10 mM) 11

2 ml 10 mM dNTPs 11

1 ml of Sigma JumpStart Taq polymerase 5U/ml 5.5

Add 70 ml of this master mix to each of the tubes. Note this 10 X PCR buffer contains 15 mM MgCl₂ leading to a final concentration of 1.5 mM Mg ⁺⁺. Finally add the following templates:

Tube

1. 30 ml of template A

2. 30 ml of template B

3. 30 ml of template C

4. 30 ml of template D

5. 30 ml of sterile water

3) Label a series of 5 sterile thin wall PCR tubes with the numbers 1-5 and your initials. Note that markings on the lid are apt to come off in the PCR machine, so you may wish to mark the side of the tubes too, and when you load and remove your samples note their position in the rack.

Note that the template is added last, and only one tube should be open at a time to minimize the possibility of template cross contamination. Because the PCR machine we use has a heated lid to prevent evaporation, it will not be necessary to overlay the tubes with mineral oil. When you have finished assembling these mixes let the TAs know, and they will transfer your samples to the PCR machine and begin cycling with the following program for 30 cycles:

94⁰C for 1 min

56⁰ C for 1.5 min

72⁰ C for 1.5 min

Figure 1 - Human b-actin Genomic Sequence Showing Internal PCR Primer Sites

This figure shows where the actin primers used in this experiment bind the human b-actin genomic DNA. PCR with these primers will produce a 288 bp product. The next diagram shows the b-actin cDNA sequence with the same primers indicated. The PCR product generated from the cDNA using these same primers is 157 bp. Why is the amplified cDNA sequence smaller than the amplified genomic sequence?

!!NA_MULTIPLE_ALIGNMENT 1.0

LOCUS HUMACCYBB 3646 bp DNA PRI 30-OCT-1994

DEFINITION Human cytoplasmic beta-actin gene, complete cds.

ACCESSION M10277

NID g177967

KEYWORDS actin; beta-cytoplasmic actin; cytoplasmic actin.

SOURCE Human DNA library from HUT-14 cell line, clone lambda-Ha160.

ORGANISM Homo sapiens

Eukaryotae; mitochondrial eukaryotes; Metazoa; Chordata;

Vertebrata; Eutheria; Primates; Catarrhini; Hominidae; Homo.

REFERENCE 1 (bases 1 to 3646)

AUTHORS Nakajima-Iijima,S., Hamada,H., Reddy,P. and Kakunaga,T.

TITLE Molecular structure of the human cytoplasmic beta-actin gene:

interspecies homology of sequences in the introns

JOURNAL Proc. Natl. Acad. Sci. U.S.A. 82 (18), 6133-6137 (1985)

MEDLINE 85298307

REFERENCE 2 (sites)

AUTHORS Harris,D.E., Warshaw,D.M. and Periasamy,M.

TITLE Nucleotide sequences of the rabbit alpha-smooth-muscle and

beta-non-muscle actin mRNAs

JOURNAL Gene 112 (2), 265-266 (1992)

MEDLINE 92210011

COMMENT A potential cap site was found at position 239. Through

interspecies sequence comparison, a second potential cap site was

found at positions 242-244.

FEATURES Location/Qualifiers

source 1. .3646

/organism="Homo sapiens"

/db_xref="taxon:9606"

/map="7pter-q22"

prim_transcript <319. .3589

/note="actin mRNA"

intron 320. .1086

/note="actin intron A"

exon <1093. .1215

/gene="ACTB"

/note="cytoplasmic beta actin, (first expressed exon);

G00-118-964"

/number=2

gene 1093. .1215

/gene="ACTB"

CDS join(1093. .1215,1348. .1587,2029. .2467,2563. .2744,

2857. .3000)

/note="cytoplasmic beta actin"

/codon_start=1

/db_xref="PID:g177968"

/translation="MDDDIAALVVDNGSGMCKAGFAGDDAPRAVFPSIVGRPRHQGVM

VGMGQKDSYVGDEAQSKRGILTLKYPIEHGIVTNWDDMEKIWHHTFYNELRVAPEEHP

VLLTEAPLNPKANREKMTQIMFETFNTPAMYVAIQAVLSLYASGRTTGIVMDSGDGVT

HTVPIYEGYALPHAILRLDLAGRDLTDYLMKILTERGYSFTTTAEREIVRDIKEKLCY

VALDFEQEMATAASSSSLEKSYELPDGQVITIGNERFRCPEALFQPSFLGMESCGIHE

TTFNSIMKCDVDIRKDLYANTVLSGGTTMYPGIADRMQKEITALAPSTMKIKIIAPPE

RKYSVWIGGSILASLSTFQQMWISKQEYDESGPSIVHRKCF"

intron 1216. .1347

/note="actin intron B"

exon 1348. .1587

/number=3

intron 1588. .2028

/note="actin intron C"

exon 2029. .2467

/number=4

intron 2468. .2562

/note="actin intron D"

exon 2563. .2744

/number=5

intron 2745. .2856

/note="actin intron E"

exon 2857. .>3000

/note="cytoplasmic beta actin"

/number=6

BASE COUNT 613 a 1117 c 1102 g 814 t

ORIGIN 145 bp upstream of AvaI site.

reformat_2.reformat MSF: 3646 Type: N January 6, 1999 13:46 Check: 5812 ..

Name: M10277 Len: 3646 Check: 5812 Weight: 1.00

1 50

M10277 GCCCAGCACC CCAAGGCGGC CAACGCCAAA ACTCTCCCTC CTCCTCTTCC

51 100

M10277 TCAATCTCGC TCTCGCTCTT TTTTTTTTTC GCAAAAGGAG GGGAGAGGGG

101 150

M10277 GTAAAAAAAT GCTGCACTGT GCGGCGAAGC CGGTGAGTGA GCGGCGCGGG

151 200

M10277 GCCAATCAGC GTGCGCCGTT CCGAAAGTTG CCTTTTATGG CTCGAGCGGC

201 250

M10277 CGCGGCGGCG CCCTATAAAA CCCAGCGGCG CGACGCGCCA CCACCGCCGA

251 300

M10277 GACCGCGTCC GCCCGCGAGC ACAGAGCCTC GCCTTTGCCG ATCCGCCGCC

301 350

M10277 CGTCCACACC CGCCGCCAGG TAAGCCCGGC CAGCCGACCG GGGCATGCGG

351 400

M10277 CCGCGGCCCT TCGCCCGTGC AGAGCCGCCG TCTGGGCCGC AGCGGGGGGC

401 450

M10277 GCATGGGGCG GAACCGGACC GCCGTGGGGG GCGCGGGAGA AGCCCCTGGG

451 500

M10277 CCTCCGGAGA TGGGGGACAC CCCACGCCAG TTCGCAGGCG CGAGGCCGCG

501 550

M10277 CTCGGGCGGG CGCGCTCCGG GGGTGCCGCT CTCGGGGCGG GGGCAACCGG

551 600

M10277 CGGGGTCTTT GTCTGAGCCG GGCTCTTGCC AATGGGGATC GCACGGTGGG

601 650

M10277 CGCGGCGTAG CCCCCGTCAG GCCCGGTGGG GGCTGGGGCG CCATGCGCGT

651 700

M10277 GCGCGCTGGT CCTTTGGGCG CTAACTGCGT GCGCGCTGGG AATTGGCGCT

701 750

M10277 AATTGCGCGT GCGCGCTGGG ACTCAATGGC GCTAATCGCG CGTGCGTTCT

751 800

M10277 GGGGCCCGGG CGCTTGCGCC ACTTCCTGCC CGAGCCGCTG GCGCCCGAGG

801 850

M10277 GTGTGGCCGC TGCGTGCGCG CGCGCGACCC GGTCGCTGTT TGAACCGGGC

851 900

M10277 GGAGGCGGGG CTGGCGCCCG GTTGGGAGGG GGTTGGGGCC TGGCTTCCTG

901 950

M10277 CCGCGCGCCG CGGGGACGCC TCCGACCAGT GTTTGCCTTT TATGGTAATA

951 1000

M10277 ACGCGGCCGG CCCGGCTTCC TTTGTCCCCA ATCTGGGCGC GCGCCGGCGC

1001 1050

M10277 CCCCTGGCGG CCTAAGGACT CGGCGCGCCG GAAGTGGCCA GGGCGGGGGC

1051 1100

M10277 GACTTCGGCT CACAGCGCGC CCGGCTATTC TCGCAGCTCA CCATGGATGA

½½½½½½½½

5’Forward Primer ATGGATGA 3’

Homologous Region

1101 1150

M10277 TGATATCGCC GCGCTCGTCG TCGACAACGG CTCCGGCATG TGCAAGGCCG

½½½½½½½½½½ ½½

5' TGATATCGCC GC 3’

Homologous Region (Continued)

1151 1200

M10277 GCTTCGCGGG CGACGATGCC CCCCGGGCCG TCTTCCCCTC CATCGTGGGG

1201 1250

M10277 CGCCCCAGGC ACCAGGTAGG GGAGCTGGCT GGGTGGGGCA GCCCCGGGAG

1251 1300

M10277 CGGGCGGGAG GCAAGGGCGC TTTCTCTGCA CAGGAGCCTC CCGGTTTCCG

1301 1350

M10277 GGGTGGGCTG CGCCCGTGCT CAGGGCTTCT TGTCCTTTCC TTCCCAGGGC

1351 1400

M10277 GTGATGGTGG GCATGGGTCA GAAGGATTCC TATGTGGGCG ACGAGGCCCA

½½½½½½½½½ ½½½½½½½½½½ ½

Reverse 3’ GTACCCAGT CTTCCTAAGG A 5’

Primer Homologous Region

1401 1450

M10277 GAGCAAGAGA GGCATCCTCA CCCTGAAGTA CCCCATCGAG CACGGCATCG

1451 1500

M10277 TCACCAACTG GGACGACATG GAGAAAATCT GGCACCACAC CTTCTACAAT

1501 1550

M10277 GAGCTGCGTG TGGCTCCCGA GGAGCACCCC GTGCTGCTGA CCGAGGCCCC

1551 1600

M10277 CCTGAACCCC AAGGCCAACC GCGAGAAGAT GACCCAGGTG AGTGGCCCGC

1601 1650

M10277 TACCTCTTCT GGTGGCCGCC TCCCTCCTTC CTGGCCTCCC GGAGCTGCGC

1651 1700

M10277 CCTTTCTCAC TGGTTCTCTC TTCTGCCGTT TTCCGTAGGA CTCTCTTCTC

1701 1750

M10277 TGACCTGAGT CTCCTTTGGA ACTCTGCAGG TTCTATTTGC TTTTTCCCAG

1751 1800

M10277 ATGAGCTCTT TTTCTGGTGT TTGTCTCTCT GACTAGGTGT CTGAGACAGT

1801 1850

M10277 GTTGTGGGTG TAGGTACTAA CACTGGCTCG TGTGACAAGG CCATGAGGCT

1851 1900

M10277 GGTGTAAAGC GGCCTTGGAG TGTGTATTAA GTAGGCGCAC AGTAGGTCTG

1901 1950

M10277 AACAGACTCC CCATCCCAAG ACCCCAGCAC ACTTAGCCGT GTTCTTTGCA

1951 2000

M10277 CTTTCTGCAT GTCCCCCGTC TGGCCTGGCT GTCCCCAGTG GCTTCCCCAG

2001 2050

M10277 TGTGACATGG TGCATCTCTG CCTTACAGAT CATGTTTGAG ACCTTCAACA

2051 2100

M10277 CCCCAGCCAT GTACGTTGCT ATCCAGGCTG TGCTATCCCT GTACGCCTCT

2101 2150

M10277 GGCCGTACCA CTGGCATCGT GATGGACTCC GGTGACGGGG TCACCCACAC

2151 2200

M10277 TGTGCCCATC TACGAGGGGT ATGCCCTCCC CCATGCCATC CTGCGTCTGG

2201 2250

M10277 ACCTGGCTGG CCGGGACCTG ACTGACTACC TCATGAAGAT CCTCACCGAG

2251 2300

M10277 CGCGGCTACA GCTTCACCAC CACGGCCGAG CGGGAAATCG TGCGTGACAT

2301 2350

M10277 TAAGGAGAAG CTGTGCTACG TCGCCCTGGA CTTCGAGCAA GAGATGGCCA

2351 2400

M10277 CGGCTGCTTC CAGCTCCTCC CTGGAGAAGA GCTACGAGCT GCCTGACGGC

2401 2450

M10277 CAGGTCATCA CCATTGGCAA TGAGCGGTTC CGCTGCCCTG AGGCACTCTT

2451 2500

M10277 CCAGCCTTCC TTCCTGGGTG AGTGGAGACT GTCTCCCGGC TCTGCCTGAC

2501 2550

M10277 ATGAGGGTTA CCCCTCGGGG CTGTGCTGTG GAAGCTAAGT CCTGCCCTCA

2551 2600

M10277 TTTCCCTCTC AGGCATGGAG TCCTGTGGCA TCCACGAAAC TACCTTCAAC

2601 2650

M10277 TCCATCATGA AGTGTGACGT GGACATCCGC AAAGACCTGT ACGCCAACAC

2651 2700

M10277 AGTGCTGTCT GGCGGCACCA CCATGTACCC TGGCATTGCC GACAGGATGC

2701 2750

M10277 AGAAGGAGAT CACTGCCCTG GCACCCAGCA CAATGAAGAT CAAGGTGGGT

2751 2800

M10277 GTCTTTCCTG CCTGAGCTGA CCTGGGCAGG TCAGCTGTGG GGTCCTGTGG

2801 2850

M10277 TGTGTGGGGA GCTGTCACAT CCAGGGTCCT CACTGCCTGT CCCCTTCCCT

2851 2900

M10277 CCTCAGATCA TTGCTCCTCC TGAGCGCAAG TACTCCGTGT GGATCGGCGG

2901 2950

M10277 CTCCATCCTG GCCTCGCTGT CCACCTTCCA GCAGATGTGG ATCAGCAAGC

2951 3000

M10277 AGGAGTATGA CGAGTCCGGC CCCTCCATCG TCCACCGCAA ATGCTTCTAG

3001 3050

M10277 GCGGACTATG ACTTAGTTGC GTTACACCCT TTCTTGACAA AACCTAACTT

3051 3100

M10277 GCGCAGAAAA CAAGATGAGA TTGGCATGGC TTTATTTGTT TTTTTTGTTT

3101 3150

M10277 TGTTTTGGTT TTTTTTTTTT TTTTGGCTTG ACTCAGGATT TAAAAACTGG

3151 3200

M10277 AACGGTGAAG GTGACAGCAG TCGGTTGGAG CGAGCATCCC CCAAAGTTCA

3201 3250

M10277 CAATGTGGCC GAGGACTTTG ATTGCATTGT TGTTTTTTTA ATAGTCATTC

3251 3300

M10277 CAAATATGAG ATGCATTGTT ACAGGAAGTC CCTTGCCATC CTAAAAGCCA

3301 3350

M10277 CCCCACTTCT CTCTAAGGAG AATGGCCCAG TCCTCTCCCA AGTCCACACA

3351 3400

M10277 GGGGAGGTGA TAGCATTGCT TTCGTGTAAA TTATGTAATG CAAAATTTTT

3401 3450

M10277 TTAATCTTCG CCTTAATACT TTTTTATTTT GTTTTATTTT GAATGATGAG

3451 3500

M10277 CCTTCGTGCC CCCCCTTCCC CCTTTTTGTC CCCCAACTTG AGATGTATGA

3501 3550

M10277 AGGCTTTTGG TCTCCCTGGG AGTGGGTGGA GGCAGCCAGG GCTTACCTGT

3551 3600

M10277 ACACTGACTT GAGACCAGTT GAATAAAAGT GCACACCTTA AAAATGAGGC

3601 3646

M10277 CAAGTGTGAC TTTGTGGTGT GGCTGGGTTG GGGGCAGCAG AGGGTG

Figure 2 - Human b-actin cDNA Sequence Showing Internal PCR Primer Sites

LOCUS HSAC07 1761 bp RNA PRI 12-SEP-1993

DEFINITION Human mRNA for beta-actin.

ACCESSION X00351 J00074 M10278

KEYWORDS actin; beta-actin.

SOURCE human.

ORGANISM Homo sapiens

Eukaryota; Animalia; Metazoa; Chordata; Vertebrata; Mammalia;

Theria; Eutheria; Primates; Haplorhini; Catarrhini; Hominidae.

REFERENCE 1 (bases 1 to 1761)

AUTHORS Ponte,P., Ng,S.Y., Engel,J., Gunning,P. and Kedes,L.

TITLE Evolutionary conservation in the untranslated regions of actin

mRNAs: DNA sequence of a human beta-actin cDNA

JOURNAL Nucleic Acids Res. 12 (3), 1687-1696 (1984)

MEDLINE 84144061

COMMENT NCBI gi: 28251

FEATURES Location/Qualifiers

source 1. .1761

/organism="Homo sapiens"

misc_feature 1. .41

/note="5'-untranslated region"

CDS 42. .1169

/note="beta-actin; NCBI gi: 28252"

/codon_start=1

/translation= "MDDDIAALVVDNGSGMCKAGFAGDDAPRAVFPSIVGRPRHQGVM

VGMGQKDSYVGDEAQSKRGILTLKYPIEHGIVTNWDDMEKIWHHTFYNELRVAPEEHP

VLLTEAPLNPKANREKMTQIMFETFNTPAMYVAIQAVLSLYASGRTTGIVMDSGDGVT

HTVPIYEGYALPHAILRLDLAGRDLTDYLMKILTERGYSFTTTAEREIVRDIKEKLCY

VALDFEQEMATAASSSSLEKSYELPDGQVITIGNERFRCPEALFQPSFLGMESCGIHE

TTFNSIMKCDVDIRKDLYANTVLSGGTTMYPGIADRMQKEITALAPSTMKIKIIAPPE

RKYSVWIGGSILASLSTFQQMWISKQEYDESGPSIVHRKCF"

misc_feature 1167. .1761

/note="3'-untranslated region"

misc_feature 1743. .1749

/note="polyadenylation signal"

polyA_site 1761

/note="polyadenylation site"

BASE COUNT 379 a 521 c 444 g 417 t

ORIGIN

X00351 Length: 1761 April 14, 1995 15:36 Type: N Check: 9510 ..

1 TTGCCGATCC GCCGCCCGTC CACACCCGCC GCCAGCTCAC CATGGATGAT

½½½½½½½½½

5’Forward Primer ATGGATGAT 3’

Homologous Region

51 GATATCGCCG CGCTCGTCGT CGACAACGGC TCCGGCATGT GCAAGGCCGG

½½½½½½½½½½ ½

5' GATATCGCCG C 3’

Homologous Region (Continued)

101 CTTCGCGGGC GACGATGCCC CCCGGGCCGT CTTCCCCTCC ATCGTGGGGC

151 GCCCCAGGCA CCAGGGCGTG ATGGTGGGCA TGGGTCAGAA GGATTCCTAT

½½½½½½½½½½½½½ ½½½½½½½½

Reverse 3’ GT ACCCAGTCTT CCTAAGGA 5’

Primer Homologous Region

201 GTGGGCGACG AGGCCCAGAG CAAGAGAGGC ATCCTCACCC TGAAGTACCC

251 CATCGAGCAC GGCATCGTCA CCAACTGGGA CGACATGGAG AAAATCTGGC

301 ACCACACCTT CTACAATGAG CTGCGTGTGG CTCCCGAGGA GCACCCCGTG

351 CTGCTGACCG AGGCCCCCCT GAACCCCAAG GCCAACCGCG AGAAGATGAC

401 CCAGATCATG TTTGAGACCT TCAACACCCC AGCCATGTAC GTTGCTATCC

451 AGGCTGTGCT ATCCCTGTAC GCCTCTGGCC GTACCACTGG CATCGTGATG

501 GACTCCGGTG ACGGGGTCAC CCACACTGTG CCCATCTACG AGGGGTATGC

551 CCTCCCCCAT GCCATCCTGC GTCTGGACCT GGCTGGCCGG GACCTGACTG

601 ACTACCTCAT GAAGATCCTC ACCGAGCGCG GCTACAGCTT CACCACCACG

651 GCCGAGCGGG AAATCGTGCG TGACATTAAG GAGAAGCTGT GCTACGTCGC

701 CCTGGACTTC GAGCAAGAGA TGGCCACGGC TGCTTCCAGC TCCTCCCTGG

751 AGAAGAGCTA CGAGCTGCCT GACGGCCAGG TCATCACCAT TGGCAATGAG

801 CGGTTCCGCT GCCCTGAGGC ACTCTTCCAG CCTTCCTTCC TGGGCATGGA

851 GTCCTGTGGC ATCCACGAAA CTACCTTCAA CTCCATCATG AAGTGTGACG

901 TGGACATCCG CAAAGACCTG TACGCCAACA CAGTGCTGTC TGGCGGCACC

951 ACCATGTACC CTGGCATTGC CGACAGGATG CAGAAGGAGA TCACTGCCCT

1001 GGCACCCAGC ACAATGAAGA TCAAGATCAT TGCTCCTCCT GAGCGCAAGT

1051 ACTCCGTGTG GATCGGCGGC TCCATCCTGG CCTCGCTGTC CACCTTCCAG

1101 CAGATGTGGA TCAGCAAGCA GGAGTATGAC GAGTCCGGCC CCTCCATCGT

1151 CCACCGCAAA TGCTTCTAGG CGGACTATGA CTTAGTTGCG TTACACCCTT

2) Using the titer determined by the class you will proceed to carry out Protocol 3.1 starting at step 4 except that we will be carrying out the procedure at ½ scale because we are using 100 mm Petri dishes which have roughly half the area of the larger plates normally used for screening libraries. Thus instead of following the exact volumes given in the protocol, carry out step 4 as follows:

Dilute the mixed phage lysate in SM buffer so that when 100 ml of Y1090 plating cells are infected with 10 ml of the diluted stock and plated, there will be ~200 - 500 plaques/100 mm plate. After you have diluted your phage, pipette 4 ml of molten top agarose from the bottle in the 55^oC water bath into a prewarmed Falcon 2063 tube in the bath and quickly return the tube to the rack in the water bath. Infect 100 ml of plating cells with 10 ml of the diluted phage for 10 min. at 37^oC. Then get prewarmed LB plates from the incubator, quickly add Y1090/ phage mixture to the top agarose, vortex and pour on the prewarmed LB plate.

This plate will be used to identify " false actin recombinants" i.e. lambda gt11 plaques carrying beta-galactosidase that react with the monoclonal anti-beta-galactosidase antibody. Remember this is a spiked “library” where we have added a far greater number of recombinant phage than would occur in a real screening. This fact allows us to plate the lysate at a much lower plaque density and use smaller plates than you would normally use in a primary screen.

3) Continue with Protocol 3.1 starting at step 4f. Today you will take the procedure through the primary antibody step given in the section on Immunoscreening Using ECL^TM, step 1. (Note: The anti- b-galactosidase antibody in this exercise is used at a 1:50,000 dilution).

Do not forget to prepare a filter for overlaying your phage library by soaking an Optitran (S&S) supported nitrocellulose filter in a solution of 10 mM IPTG (an inducer of the lac operon) for 10 min., then place the filter on a piece of Whatman 3MM paper and leave the filter to dry at room temperature. Keep the filter clean by covering it with aluminum foil or a clean paper towel until you need it later this afternoon.

Be sure to watch the plates in the 42^oC incubator and to put the IPTG soaked filter on the plates at 3¹/₂ hours from the time of plating. After applying the filter, downshift your plate to 37^oC for a second 3 to 3¹/₂ h incubation. This will give plenty of time for the phage to lyse and for the filters to pick up the expressed "false actin fusion protein" i.e. beta-galactosidase.

Before removing the filter after the 37^oC incubation be sure to mark the position of your filters in their Petri dishes by sticking them with a hypodermic needle dipped in ink (if you have not already done so) to aid in subsequently aligning your plates with the film and carry out plaque lifts as described in Protocol 3.1. This technique will be demonstrated for you.

Make sure you understand why you are using a filter impregnated with IPTG and why the plates being incubated initially at 42^oC rather than the usual 37^oC in this protocol.

Phagemid Rescue (continued)

1) You will next need to grow the recombinant b-actin phage you cored yesterday so that you can rescue the phagemid by superinfection. You will carry out the rescue as described in Protocol 3.5. A diagram of the excision process is given in the protocol.

2) At step 11 of Protocol 3.5, infect only one tube of SOLR cells (200 ml) with 10 ml of phage stock. We have found that the rescue procedure is so effective that 10 ml of the crude phage stock will give you plenty of ampicillin resistant colonies.

3) At step 13, plate 1 and 100 ml of your infected SOLR cells on LB plates containing ampicillin, and place the plates at 37^oC overnight. When you are plating small volumes of cells between 1 to 10 ml, you should first spot your plate with 100 ml of LB broth. Then add the 1 to 10 ml of cells to the middle of the LB spot and spread. This makes it easier to spread the volume of cells uniformly on the plates.

Identification of Recombinant Vectors Carrying Inserts Using the LacZ System

1) The TAs will demonstrate how to pour a top agar plate that allows for discrimination of Lac + phage (lacking an insert) from Lac- phage (carrying an insert) by means of a color reaction catalyzed by b-galactosidase. The phage mixture being used in this demonstration is the same stock of lambda wild type lacking an insert and lambda gt11 carrying a b-galactosidase insert that was for the antibody screening experiment. While we will not really use this technique in our course, it is a powerful method for identifying clones containing inserts into the b-galactosidase gene that is widely used in molecular biology.

Blackboard Notes Day 4 (Thur)

READ ALL OF THIS SHEET BEFORE PROCEEDING

Today you will be developing your plaque lifts that are currently incubating with primary antibody to identify b-galactosidase positive "false actin" recombinant phage. Then you will continue the phagemid rescue of actin-containing phage lambda plaques. Next you will examine plates prepared by the TAs containing a mixed lysate of the Lac⁺ and Lac^- phage which demonstrate how using the LacZ system vectors carrying inserts can be distinguished from non-recombinant vectors. Finally you will start preparing DNA from your rescued b-actin clone in the Bluescript phagemid which will ultimately be subcloned into an E. coli expression vector.

cDNA Screening (Continued)

1) Begin by washing your filters 2 X for 10 min. in ~ 100 -150 ml cold PBST to remove the unbound primary mouse monoclonal anti b-galactosidase antibody. Next we will provide you with the secondary anti-mouse IgG antibody linked to horseradish peroxidase (HRP) diluted at 1:30,000 in PBST 5% nonfat dry milk. You will then incubate this secondary antibody with your filters for 30 minutes just as you did for Western blotting except that this time you will be developing your filters in the Pierce Super Signal substrate to identify antibody positive plaques which express beta galactosidase i.e. “false actin”. Western blots can in theory be carried out with either an alkaline phosphatase or HRP reporter enzyme. Our choice of which reporter to use here was determined by the supported nitrocellulose membrane being used. Nitrocellulose membranes quench the signal from alkaline phosphatase reaction unless specific modifiers are used. This leads to a lower signal than with HRP.

Phagemid Rescue (Continued)

1) You will also need to prepare two test tubes containing 5 ml LB containing 50 mg/ml ampicillin. You will be provided with a 50 mg/ml stock of ampicillin. Inoculate the two tubes with separate ampicillin resistant colonies from your phagemid rescue experiment. The rescue results in generation of pBluescript SK- phagemid that contain an ampicillin resistant gene for selection as well as the b-actin insert. Place this tube in the 37^oC shaking incubator so that it is well aerated. This culture will be used tomorrow to prepare plasmid DNA for subcloning into an expression vector that provides very tight regulation of gene expression. This vector will allow us to express b-actin in E. coli while minimizing some of the problems that result from attempting to produce the toxic b-actin protein from the leaky b-galactosidase promoter in the Bluescript plasmids.

Identification Of Recombinant Vectors Carrying Inserts Using The LacZ System (Continued)

1) Examine the plate of mixed lambda wild type (Lac^-) - lambda gt11 (Lac⁺) that the TAs plated out yesterday with IPTG and XGAL. You should see both white and blue plaques. Make sure you understand why Lac⁺ plaques are blue and Lac^- plaques are white. You should also understand how the DNA fragment containing the a-peptide of b-galactosidase/multiple cloning region construct can be cloned into other vectors and plasmids and why that would be useful. We will not be able to use blue/white identification in our own subcloning experiment into plasmid pET28b since this vector does not contain the a-peptide of b-galactosidase.

Demonstration

1) How to pour an agarose gel.

Blackboard Notes Day 5 (Fri)

READ ALL OF THIS SHEET BEFORE PROCEEDING

Today will be devoted to preparing high quality plasmid DNA for subcloning and automated DNA sequencing (We will not actually be carrying out automated sequencing in the course, but will introduce you the technique and show you how to submit samples to the UNC facility). You will also initiate the subcloning of the b-actin insert into a bacterial expression plasmid, and begin the isolation of chromosomal DNA you will need for Southern blotting.

Prepare Plasmid DNA For Subcloning

(This DNA could also be used for sequencing as well)

1) First you will prepare plasmid minipreps from the pBluescript SK-/b-actin cultures you inoculated last night using a protocol that exploits the ability of a proprietary ion exchange resin to separate DNA from RNA and protein contaminants (see Protocol 4.5). You will use two of the Qiagen Tip-20s to yield enough DNA for the subsequent subcloning procedure. If this procedure seems like magic to you, read Protocol 4.6, which tries to explain the initial steps of the procedure.

2) Next quantitate the recovery of plasmid DNA using Protocol 4.1. We will demonstrate the use of the fluorometer. Record the concentration. Three mg of this DNA will be used for the subcloning procedure described below. One microliter of the remaining DNA can be diluted to about 1 ng/ml in sterile TE. Mark the tube and store it at 4^oC where you will remember where you put it. This DNA will be used for one of the templates used in the PCR experiment done on day 7.

Demonstration

1) How to quantify DNA using a fluorometer

2) How to quantify DNA using a spectrophotometer

Subcloning

1) Next you will isolate the b-actin insert DNA from the plasmid you just prepared for subsequent subcloning into a bacterial expression vector pET28b. The construction of this vector will be discussed in class. You should also look at the diagrams of the pET28 vectors on the following page.

To carry out this cloning, both the pBluescript plasmid containing the insert and the pET28b vector into which the insert will be cloned have to be cut with the restriction enzymes Eco RI and Xho I. Conveniently, both of these enzymes use the same digestion buffer. (How would you deal with the situation when this was not the case?) You may wish to check one of the catalogs for some information about these enzymes and Protocol 9.1 for some general information about restriction enzyme digests. The restriction enzymes and the buffer will be provided to you. You will be provided with the doubly cut pET28b vector, but you will have to prepare the insert yourself.

Each of you should digest 5 mg of the plasmid DNA prepared in step 1 and quantitated in step 2. You should set up your restriction enzyme digestion as shown below and allow it to go for 1 hour at 37^oC .

Question: Assuming complete digestion of the DNA with the restriction enzymes and 50% recovery of the insert DNA after the agarose gel purification procedure, about how much insert DNA do you expect to recover?

5 mg pBluescript/b-actin plasmid x ml

Water 24-x ml

10x Reaction Buffer H 3 ml

Eco RI (10 U/ml) 1.5 ml

Xho I (10 U/ml) 1.5 ml

Total 30 ml

Mix the tubes gently by pipetting and/or finger flicking followed by spinning for a few seconds in the microcentrifuge to ensure that all the components are mixed. Note that in this particular digestion, simultaneous digestion with the two enzymes works well.

Question: What might happen if you needed to digest with the enzymes SpeI and BamHI (see page 312 of the Stratagene manual).

Question: How many fold over digestion with the enzymes Eco RI and Xho I have you used in this reaction?

2) While your DNA is digesting, you need to prepare an 0.8% agarose gel for purification of the b-actin DNA insert. Because the standard wells on our gel combs will only hold ~ 10 ml, you will need to modify the gel comb so as to hold the 30 ml digestion. This can easily be done by covering two adjacent teeth on the comb with gel sequencing tape. We will demonstrate how to do this. After you have taped your combs, we will provide you with a solution of 0.8% agarose in 0.5X TAE buffer. The agarose has been premelted in a microwave oven and allowed to cool to about 55^oC and is ready to pour. Note: The bottle will be very warm, but not scalding hot. Remove the bottle containing the agarose and pour the gel quickly. Immediately return the agarose to the water bath to prevent the agarose from solidifying.

3) After the gel has solidified, pour enough buffer into the gel box so that it will just cover the gel. Gently remove the comb and attach the leads to the power supply and make sure the gel box is functioning. Prepare the following samples for loading on the gel:

Sample TE 10X Loading Final

Vol (ml) (ml) Dye (ml) Volume (ml)

1) (500 ng) of DNA 1 kb ladder from Sigma 5 4 1.0 10

2) 0.25 mg of your uncut plasmid DNA X 9-X 1.0 10

3) 100 ng of cut insert which will serve as a 9 0 1.0 10

marker where you expect to find your insert

4) restriction enzyme digestion of 30 0 3.3 33.3

pBluescript/b-actin

Both the cut insert and size markers will be provided. Load your gels in the following order:

Lane 1 10 ml of 1 kb ladder DNA markers (500 ng)

Lane 2 10 ml of uncut plasmid (0.25 mg)

Lane 3 10 ml of cut insert (100 ng)

Lane 4 blank

Lane 5-6 (taped) 33 ml (3 mg) of restriction enzyme digest

Note that we will not heat inactivate the restriction enzymes as it does not appear necessary in this case. If you were preparing the vector DNA yourself, you would also load an aliquot of the digested vector to see what fraction remained undigested. Too much undigested vector (more than a few percent) can make identification of recombinant colonies difficult. If the digestion of the vector was not complete, you would have to run the reaction on a preparative gel and purify the cut vector away from the undigested DNA prior to setting up the ligations.

Electrophoresis at 50 V for 1-2 h until the bromophenol dye is about 2/3 to 3/4 the way down the gel. Turn off the power. Place the gel into the ethidium bromide staining solution provided (wear gloves!). Stain for about 10 minutes then photograph your gel. You should try to minimize the exposure to UV to reduce the chance of mutating the DNA. The band you will isolate is the one that has moved furthest down the gel and should be opposite the marker in lane 3. A good practice is to have a scalpel ready before illuminating the gel and to quickly make a cut above and below the band as rapidly as possible. Then you can turn off the light and finish cutting out the band. You should try to take only the band and leave any extraneous agarose behind. Carrying excess agarose along to the purification step will lower DNA recovery. Store the gel slice overnight at 4^oC in a preweighed microfuge tube.

Southern Blotting

1) Start the preparation of chromosomal DNA from HeLa cells for your Southern blot following Protocol 4.4. You will be provided with 30 ml of HeLa cells at a concentration of 5 x 10⁵ cells/ml. After pelleting the cells, pour off the supernatant. Continue with the protocol through Step 5, which is an overnight incubation with Proteinase K.

Sequencing

1) We will have a lecture /movie explaining the principles of automated DNA sequencing.

Movie

As time allows we will watch the video "The Race For The Double Helix".

Blackboard Notes Day 6 (Sat)

READ ALL OF THIS SHEET BEFORE PROCEEDING

Today you will recover your b-actin insert from the agarose gel slice, estimate its concentration, and set up a ligation of the insert into the pET28b expression vector, along with the appropriate controls. Then you will continue your chromosomal

DNA preparation needed for Southern blotting.

Subcloning (Continued)

1) At this point you will purify the insert from the gel slice following Protocol 4.25. You will need to weigh your microcentrifuge tube before and after you put your gel slice in it to determine the gel slice weight. Assume the density of the gel slice is ~1 g/ml.

2) After purifying the insert from the gel slice, quantitate the amount of insert by fluorometry - Protocol 4.1. The DNA will be far too dilute to detect by spectrophotometry unless you use a microcuvette.

3) Before you go home you will need to set up the ligation of your 1.4 kb XhoI-EcoRI fragment into the Novagen expression vector pET28b. We will discuss the elegant design of this vector later. For now you need to read carefully the discussion of ligation on page 6.1 and especially the ligation controls on page 6.2. The ligation protocol itself is Protocol 6.1 A. We will provide you with purified insert and uncut and doubly cut pET vector. These will be used along with your purified insert as controls for the control ligations discussed below (following page).

Now set up the following 20 ml ligations:

	Vector	Insert	Water	10X Ligase Buffer	Ligase
1. Sterile Control*	-		17 ml	2 ml	l ml (0.1 U)
2. Competence Control**	200 ng Uncut Vector in 2 ml	-	16 ml	4 ml	-
3. Cut Vector Control	200 ng Doubly Cut Vector in 2 ml	-	16 ml	2 ml	-
4. Cut Vector Control With Ligase	200 ng Doubly Cut Vector in 2 ml	-	15 ml	2 ml	l ml (0.1 U)
5. Complete Ligation With Provided Insert	200 ng Doubly Cut Vector in 2 ml	200 ng in 5 ml	10 ml	2 ml	l ml (0.1 U)
6. Complete Ligation with Your Insert	200 ng Doubly Cut Vector in 2 ml	200 ng in X ml	15-X ml	2 ml	l ml (0.1 U)

* This not a perfect sterile control, but at least it insures everything except the purified insert and vector are sterile.

Four further controls could be added to this already formidable list. These are singly cut vector with each of the two enzymes, plus and minus ligase in order to test that the ligase is active. We have already determined that our ligase is active, so we can omit these controls. However tomorrow the TAs will transform with 200 pg of uncut vector DNA to check the maximum efficiency of the cells, so you can see how much less efficient transformation is at the 20 ng level that we use when we plate 1/10 of our ligation mixes.

You will be provided with a tube containing 10X Ligase Buffer and another with sterile water. When you are ready for it, the ligase will be dispensed from the front desk. Incubate all tubes O/N at 16^oC in a heat block in the cold room.

Southern Blotting (Continued)

1) You need to complete your genomic prep today - Protocol 4.4 starting at step 6. In the final step, when you attempt to dissolve the DNA in the TE, put the tube at 37^oC overnight to be sure the DNA dissolves completely. Do not vortex the DNA to hasten dissolution as this might shear the DNA to an unacceptably low MW.

Blackboard Notes Day 7 (Sun)

READ ALL OF THESE NOTES BEFORE PROCEEDING

XX It's probably better to do the transformations first, so that the cells will have maximum time to grow O/N. The first thing you should do today is to determine the concentration of the chromosomal DNA preparation you have been making for the Southern Blot. Then you will set up PCR reactions to amplify the b- actin gene from various templates . The templates will include purified plasmid DNA containing a human b-actin c-DNA insert, a crude preparation of plasmid DNA carrying the same insert , purified HeLa genomic DNA that you will use later for the Southern blotting, and a crude preparation of HeLa genomic DNA. Finally you will transform NovaBlue competent cells with your b-actin insert-pET28b ligation mix and the controls

Southern Blotting (Continued)

1. Use the fluorometer to determine the concentration of your genomic DNA (Protocol 4.1). If time allows also determine the DNA concentration by spectrophotometry (Protocol 4.2). How do the two results compare? How can you explain any difference? Note: You will probably find that a good fraction of your DNA fails to go into solution even after additional heating. You may wish in this case to add a bit more TE buffer and try heating the solution again. Try however NOT to dilute the DNA too much. We want to aim for a final DNA concentration of about 0.5 ug/ul so that the DNA will be concentrated enough to avoid problems in setting up your restriction enzyme digestions tomorrow

Subcloning (Continued)

1. In the afternoon you will transform 20 ml aliquots of competent NovaBlue cells following Protocol 6.4, Procedure A and Common Steps A, B, and C. These competent cells were obtained commercially from Novagen, but we provide protocols for making your own competent cells in the manual. Thaw the cells on ice as indicated in the protocol. You may find a considerable volume of cells in the lid of the vial. If so, tap the vial so that all the liquid collects in the vial or spin for a second in a microfuge to collect this material. In any case resuspend the cells GENTLY by pipetting up and down. You will transform plates from ligation conditions 1-6 in the ligation chart given in yesterday's instructions. For tubes 1, 3-6 take 10 ml of the final 100 ml transformation mix and add it to 100 ml of LB prespotted on the middle of an LB Kan plate. Then spread the mixture. For tube 2 (competence control), dilute an aliquot of the transformation mix 1:10 in LB. Then take 10 ml of the resulting dilution and plate as above.

Note: Other non-expressing strains such as BL21 or XL-1 Blue can be used at this step as well, but be sure to note the efficiency of the strains before setting up your transformation reactions.

Question: Why are cells lacking the DE3 marker unable to express the construct?

Dinner at 6 PM tonight is at Phil’s house 2508 Foxwood Drive in Chapel Hill. Telephone is 967-3530.

Blackboard Notes Day 8 (Mon)

READ ALL OF THESE NOTES BEFORE PROCEEDING

Today you will examine the transformation plates from yesterday to see if the transformation proceeded as expected. If the colonies are very small, leave them in the incubator and proceed to the running the PCR products from yesterday's amplification on an agarose gel (see below). After you have tallied the number of colonies on each plate, you will identify colonies containing b-actin inserts using Protocol 4.9. Next you will set up digestions of your HeLa chromosomal DNA for Southern blotting. Following this you will run aliquots of the PCR reactions that you set up yesterday on an agarose gel to see if you were successful in amplifying the b-actin DNA from the various templates. Finally you will set up a reaction to prepare DIG labeled random primed b-actin DNA which you will later use to probe your Southern blot.

Subcloning (Continued)

1) The first thing you should do today is to examine your transformation plates. Calculate the efficiency of the competent cells from transformation tube 2 which contained uncut vector. Express your results as colonies/mg uncut plasmid DNA. The claimed efficiency of the NovaBlue cells is 10⁸ colonies/mg of uncut plasmid. How does your result compare to this? How might you explain any discrepancies? You should also be able to calculate what fraction of the DNA remained uncut after the Eco RI and Xho I digestions. This fraction can be determined from the number of colonies seen on plate 2 assuming that all the colonies represent transformation by uncut vector. This value will probably be in the range of 0.1 to 1%. Examine the plate of doubly cut vector lacking insert that has been religated (Plate 4). Does it show an increase in colonies compared to the unligated doubly cut vector? If so, how would you explain this? Finally examine your plates transformed with your insert and ours. One would hope to see that about 5-30% of the vectors ligated to the insert.

2) Because we have cloned into a vector that lacks the convenience of the blue/white colony selection feature, we have no easy way of knowing which of our colonies actually has an insert in the pET plasmid. Therefore we will need to screen multiple clones from your recombinant plates to see which ones have an insert. This can be done by any of the multiple miniprep methods for preparing plasmid DNA or by PCR methods, but the cheapest and fastest way is to use Protocol 4.9. In this method we make very crude lysates of total cells and run them on gels. Be sure to transfer each colony to the master plate containing LB Kan as described in the protocol. You have grids on page 2-17 of the manual which you can use to guide you in making the plate. Incubate the master plate at 37^oC O/N to grow the patched transformants. This plate will be used tomorrow to retrieve colonies containing recombinant plasmids as determined by the Direct Lysis Protocol (Protocol 4.9). To carry out the Direct Lysis Protocol, you should make a 0.8% TBE agarose midigel that contains 0.05% SDS. Use approximately 75 ml of the agarose. If the gel is too thick, it will take too long to stain and it will be difficult to see the DNA. Use two 16 well combs at the top and middle of the gel as noted in the protocol. Note that this is not the standard TBE buffer used for DNA electrophoresis because it also contains SDS to help prevent the proteins in the crude lysate from binding to the DNA. We will use two combs and run the gels only until the bromophenol blue marker of the upper part of the gel reaches the lower comb. You should load your gel as follows:

Lanes 1 and 16 - Crude lysate from non-recombinant bacteria obtained from your plate transformed with uncut plasmid. Note: If these colonies are too small you can pool several since they should all be the same.

Lanes 2-15 Crude lysate made from colonies on your potential recombinant plates. Note: If these colonies are small, let them grow for a few more hours before performing this procedure because small colonies contain insufficient DNA to give a good signal on the gel.

Each student should load 14 potential recombinants by sharing a gel with his/her partner. Each pair will thus share a single double-combed midigel.

Because we will be loading lysates into wells in both the upper and bottom half of the gel, you only be able to run the gels for about 1 h, which will be sufficient for our purposes, compared to the 2-2¹/₂ hours mentioned in the protocol. Following electrophoresis you should stain your gel in ethidium bromide (Protocol 4.19). You will have to leave the gel in the stain for about an hour to see good staining. You should look for any plasmids that move slightly more slowly than your non-recombinant plasmid control. These are your potential recombinants. You may also see some plasmids that migrate more rapidly than your PET28b control. How might you account for these? You may also see a band corresponding to the pLysS plasmid.

3) Incubate the master plate overnight at 37^o C.

4) If time allows pour a 0.8% minigel and run out the remaining ligation mixture (9 ml plus 1 ml loading dye) in order to observe the effects of ligation on the molecular weight of the DNA. Load 10 ml of Boeringher Mannheim Marker II DNA size markers on the gel as well. Run the gel at 50 V until the bromophenol blue is at least 2/3 the way down the gel. A list of the fragment sizes will be posted at the front of the room. This gel should allow you to visualize the result of your DNA ligations since ligated samples will form higher molecular weight products.

PCR (Continued)

1) Pour a 4% NuSeive 3:1/ 0.5X TAE agarose minigel and run the PCR products from yesterday's PCR experiment. Use 10 ml from each reaction + 2 ml of 6X loading dye and include one lane of 5 ml of 100 bp markers at 50 ng/ml (provided) plus 5 ml of TE. If time allows try to run the gel at 50 volts until the orange marker is about at the bottom of the gel. This will take about 2 1/2 hours. You can run the gel at 100 volts but the resolution will probably be poorer. NuSeive 3:1 agarose is a standard melting temperature agarose especially developed for resolution of DNA fragments up to 1000bp.

2) Stain your gel with ethidium bromide and photograph. Determine the size of the band(s). Do you see a 288 bp band in any of your HeLa genomic samples? Do you see a 157 bp cDNA band in your bacterial plasmid templates. Are there any other bands in the genomic template? If so, what are their sizes? Are your results consistent with there being cDNA-like sequences in HeLa genomic DNA?

Southern Blotting (Continued)

1) You will next set up the overnight enzyme digests (Protocol 4.18) for the Southern according to the chart below. In this chart Tube 1 is the Eco RI digestion of your genomic DNA preparation. Tube 2 is a control for any nucleases contaminating your genomic DNA which are active in H buffer. Tube 3 is the Hind III digestion in B Buffer. Tube 4 is a control for any nucleases contaminating your DNA which are active in B buffer. Tube 5 is a control containing 10 mg genomic DNA and 400 pg of a plasmid containing a b-actin insert (equivalent to 100 pg b-actin). This controls for both completeness of the restriction enzyme digestion and serves as a positive control for transfer and hybridization. Tube 6 is the Eco RI digestion of the provided genomic DNA preparation.

	10X Reaction Buffer	Water	Genomic DNA	Plasmid DNA (provided)	Eco RI	Hind III
Tube 1	3.0 ml H buffer	24-X ml	X ml (10 mg)		3.0 ml	----
Tube 2	3.0 ml H Buffer	27-X ml	X ml (10 mg)		----	----
Tube 3	3.0 ml B Buffer	24-X ml	X ml (10 mg)		----	3.0 ml
Tube 4	3.0 ml B Buffer	27-X ml	X ml (10 mg)		----	----
Tube 5	3.0 ml H buffer	23-X ml	X ml (10 mg)	1 ml (300 pg)	3.0 ml	----
Tube 6	3.0 ml H buffer	24-X ml	X ml (10 mg (provided)		3.0 ml	----

Making A Random Primed Probe For Southern Hybridization

1. Each pair of students should start making a random primed human b-actin probe for the Southern blot according to the Genius Kit Random Primed DNA Labeling Procedure (Protocol 5.9). We will use 120 ng of b-actin insert DNA that has been gel purified per 20 ml reaction. We will provide this to you. Incubate your labeling reaction overnight at 37^oC. You should obtain up to 280 ng of labeled probe during a 20 h reaction according to the Genius Protocol Book. In addition, one person per bay should label the control DNA supplied with the Genius Kit. Should our DNA fail to label well, this control will help us distinguish between problems with the kit components and problems with our DNA.

Blackboard Notes Day 9 (Tue)

READ ALL OF THIS SHEET BEFORE PROCEEDING

The first thing you will do today is to set up an experiment to determine the optimum conditions for amplifying b-actin DNA from either purified or relatively crude human genomic DNA. You will next check your restriction enzyme digestions of your chromosomal DNA for Southern Blotting on an agarose minigel to be sure they cut to completion and that there was no digestion of your DNA by contaminating DNases. If the results indicate that the cutting was good, you will next pour and load an agarose midigel which will be subsequently Southern blotted tomorrow. You will also assess the quality of the random primed digoxigenin labeled b-actin probe you made yesterday for the Southern hybridization procedure. The last thing you will do is to inoculate two 5 ml cultures with colonies from your master plate containing pET28B/b-actin inserts as determined by the rapid screening protocol you carried out yesterday. These cultures will be used to prepare plasmid DNA for transforming BL21(DE3) pLysS cells for expression of human b-actin in E. coli.

PCR Optimization

1) The purpose of this exercise is to demonstrate the need for optimization in PCR reactions. You will be using the Strategene Opti-Prime optimization kit - Protocol 8.7. In this case you will determine the optimal pH and Mg⁺⁺ concentrations needed to make the PCR reaction work using about 200 ng of purified of relatively crude human genomic DNA. One person per bay should dilute a small aliquot of the genomic DNA you are preparing for the Southern blot to 100 ng/ml in 50 ml water. Each bay will be provided with the 12 Opti-Prime buffers and the b-actin forward and reverse primers (10 mM) and one person will make a master mix for 13 100 ml reactions as follows:

Per 1X Per 13

reaction (100 ml) reactions (100 ml)

Water 81 1053

b-actin forward primer (10 mM) 2 26

b-actin reverse primer (10 mM) 2 26

Genomic DNA (100 ng/ml) 2 26

10 mM dNTPs 2 26

Once the master mix is made, aliquot 89 ml into 12 provided PCR tubes and give each person in the bay three of the tubes. Each tube already contains 10 ml of one of the Opti-Prime 10X Buffers. Finally add 1 ml of polymerase and proceed with the PCR reaction. A tube of polymerase for each bay will be provided by the TAs. The polymerase we will be using is the Sigma Jump Start version of Taq which allows one to carry out an easy hot start procedure.. Place all the tubes from a given group together in the PCR machine. It is difficult to label these tubes so you need to identify your samples by their position in the grid that is printed on the block that holds the samples.

2) We will use the following program for the PCR

30 cycles 1 min 94^oC to denature sample

1.5 min 56^oC to anneal primer

1.5 min 72^oC extension

3) Pour a 4% NuSeive 3:1/ 0.5X TAE agarose minigel and run the PCR products from your optimization experiment. Use 10 ml from each reaction + 2 ml of 6X loading dye and include one lane of 5 ml of 100 bp markers at 50 ng/ml (provided) plus 5 ul of TE. If time allows try to run the gel at 50 volts until the orange marker is about at the bottom of the gel. This will take about 2 1/2 hours. You can run the gel at 100 volts but the resolution will probably be poorer.

Southern Blotting (Continued)

1) Each student should pour a 0.8% 0.5 x TAE minigel and check your overnight digestions of HeLa chromosomal DNA. Dilute 5 ml from each of the four digest tubes from step 4 on Sunday with 4 ml of TE and add 1 ml of 10X tracking dye.

Each student should load

Lane 2 Chromosomal Digest with Eco RI from Tube 1.

Lane 3 Chromosomal DNA incubated with H buffer alone (control for nuclease contamination) from Tube 2.

Lane 4 Chromosomal Digest with Hind III from Tube 3.

Lane 5 Chromosomal DNA incubated with B buffer alone (control for nuclease contamination) from Tube 4.

Lane 6 EcoRI digest of chromosomal DNA prepared by TAs from tube 6.

Lane 7 10 ml (500 ng) of Lambda Hind III DNA molecular weight markers.

Run this gel for about 1 h at 100 V, stain with ethidium bromide and take a picture. You should see clear evidence for cleavage of the DNA only in tubes containing the restriction enzymes.

2) In the afternoon, each pair of people should pour a 0.5X TAE midigel for running the restriction enzyme digest which will subsequently be Southern blotted. For this experiment the TAs will have already added 0.8 g of SeaKem GTG agarose (BioWhitter - formally FMC) to 100 ml of room temperature 0.5X TAE electrophoresis buffer (a 0.8% gel) and heated the solution until the agarose has dissolved. In several instances the volumes we will need to load will be greater than one well can contain, so we will again have to create double wells using sequencing tape as we did on Day 5.

Student 1 will load the following:

Lane 1 is a single well and will contain 500 ng lambda Hind III molecular weight markers in 10 ml.

Lane 2 will be a double well containing the remaining 9.5 mg of your Eco R1 digest of HeLa DNA in 30 ml.

Lane 3 will be a double well containing the remaining 9.5 mg of the Hind III digest of HeLa DNA in 30 ml.

Lane 4 will be a single lane containing 1 pg of linearized pBluescript human b-actin in 10 ml containing 10 mg salmon sperm DNA.

Lane 5 will be a single lane containing 10 pg of linearized pBluescript human b-actin in 10 ml containing 10 mg salmon sperm DNA.

Lane 6 will be a single lane containing 30 pg of linearized human b- pBluescript actin in 10 ml containing 10 mg of salmon sperm DNA.

Note that normally one would use your human genomic DNA in place of the salmon sperm DNA spiked with linearized plasmid containing the b-actin insert as a positive control. This control would have allowed you to test the possibility that an inhibitor in your genomic DNA preparation was preventing the DNA from digesting since such an inhibitor would presumably prevent the digestion of the plasmid DNA as well. If the plasmid remained undigested you would know this because you would see the multiple forms of the plasmid on the Southern Blot rather than the single band expected for a completely cut plasmid. Herring or Salmon sperm DNA containing linearized plasmid was used in this instance to reduce the need for each student to isolate and digest large amounts of genomic DNA and because we wanted to provide you with a positive control. Using the same amount of DNA in the control lanes as in the human DNA digests gives more reliable results. Many times small amounts of linerarized plasmid will not bind the membrane as efficiently when it is loaded by itself as it does when transferred along with the other DNA.

Student 2 will load in the same order lanes 7-12.

The plasmid controls have been included to determine the limits of detection of your probes. Run the gel at 25 V overnight. Load the gel just before you go home.

6. Next you will assess the success of your random primed labeling of the b-actin probe you made last night by making probe test strips in which you will spot varying amounts of your probe on a provided membrane strip. You will also be provided with a membrane strip that we have prespotted with dig-labeled DNA purchased from Roche. Both strips will then be developed using the colorimetric assay that comes with the Genius kit and is described in Protocol 5.9 starting at step 6.

When you have completed spotting and developing your random primed test strip, you should make a second test strip of denatured target b-actin DNA which will be provided. This test strip of unlabeled DNA will be included in the hybridization bottle along with your Southern blot. (Why is it important that this DNA be denatured?) You should dilute and spot this target DNA and a buffer control just as you did the probe DNA, so that the final amounts spotted on the filter range from 100 pg to 0.1 pg. You need not spot any control DNA however as you did in making your probe test strip. The dilution series on this test strip will allow you to determine the sensitivity of your probe under real hybridization conditions independent of any questions about the efficiency of DNA transfer. After making this strip and UV crosslinking in the Stratalinker, you should wrap the target test strip in Saran Wrap and place it in your fridge until time to include it in the prehybridization and the hybridization steps along with the Southern blot.

Subcloning (Continued)

1) Before you go home, inoculate 2 or 3 of your recombinants from your master plate clones into 5mLs of LB^kan Broth so that tomorrow you will be able to make minipreps of the plasmid DNA. This DNA will then be used for transforming BL21(DE3) pLysS cells for expression.

2) 2) Store your expression master plate from yesterday at 4^o C.

Blackboard Notes Day 10 (Wed)

READ ALL OF THIS SHEET BEFORE PROCEEDING

Today you will carry out the Southern transfer of your digested HeLa chromosomal DNA and fix the DNA to a filter with UV light. Next you will set up synthesis of a b-actin riboprobe and assess the quality of the probe which will then be stored frozen at -70⁰C. overnight. Then you will use the remainder of your 20 ml random primed probe synthesis to set up an overnight hybridization with the human b-actin gene on the Southern blot. Finally you will carry out a rapid plasmid preparation on the two cultures of b-actin recombinant plasmids, determine the recovered DNA concentrations, and use that DNA to transform BL21 (DE3) pLys S for subsequent b-actin expression.

Southern Blotting (Continued)

1) Each group of two people should stain and photograph their gel of the restriction enzyme digest (Protocol 9.2). Minimize exposure of the gel to UV light and be careful not to drop the slippery gels. We will be using the alkaline blotting method described in Protocol 9.3. Begin by treating the gel with NaOH for 15 minutes in a clean plastic tray to denature the DNA as you set up a downward transfer for 1.5 to 2 hours as described in the protocol. This will be demonstrated in class.

2) After 1.5 to 2 hours, take down the Southern downward transfer and process the filter according to Protocol 9.3. Before placing the blot in the hybridization bottle, cut the membrane in two between lanes 6 and 7 with clean scissors so that each student will be responsible for hybridizing his or her own lanes. In addition, cut off an inch from the bottom of the blot so that it fits into the bottle better. Be sure that the blot is flush against the glass bottle and that the DNA side is up so that even hybridization will occur. We will use 10 ml of Dig-EasyHyb prehybridization solution in the short Bellco bottles, and carry out prehybridization at 42^oC. Be sure to include the target b-actin dilution series target strip which you prepared yesterday and make sure the strip is placed in the bottle so that it is face up against the side of the glass bottle as well.

3) Just before the end of the prehybridization step each student should remove 5 ml of your random primed b-actin probe, or enough of the probe to give 10 - 15 ng/ml in the final 10 ml of hybridization solution and add it to 100 ml TE. Immediately boil for 10 minutes then snap cool in an ice water bath for 5 -10 min.

Note: The hybridization solution used in this procedure contains a proprietary reagent (urea?) which greatly lowers the temperature at which DNA strands separate. The concentration of this reagent is such that the hybridization conditions are analogous to using 50% formamide solution. When using a non-formamide based hybridization solution, the first time you use a probe after it is synthesized you must boil the DNA for 5 to 10 minutes and snap cool in an ice water bath to assure the DNA strands are denatured. The denatured probe can then be added to the hybridization solution. In subsequent uses of the probe, which we do not do here, it is important to heat the hybridization solution containing the used probe to a temperature sufficient (65-68^oC) to denature any duplex that formed during the first use of the probe.

Next add the denatured probe to 10 ml of Dig EasyHyb hybridization solution in a 15 ml tube that has been preheated to 42^oC. Preheating the solution presumably increases specificity and helps reduce background. Use this probe within 30 minutes to avoid extensive reannealing of the probe. This would reduce the effective single strand concentration and hence reduce the sensitivity of the hybridization.

4) Remove the prehybridization solution and immediately add your denatured probe in hybridization solution. Incubate your hybridizations overnight at 42^oC.

Northern Blotting

1) Each group of four people will make a riboprobe as described in Protocol 5.11, steps 1-4. You will be provided with 1 mg of the linearized pBluescript-b-actin plasmid which you will transcribe. Because we know the orientation of the insert in this vector, we know that if you use the T7 promoter site and T7 RNA polymerase, you will transcribe the antisense strand which will be complementary to the b-actin mRNA (see notes on transcription below).

Questions: How would you make a probe of the sense strand if you wished to do so? Suppose you didn’t know the orientation of the insert in this plasmid. Can you suggest a way the orientation could be determined without sequencing the plasmid?

Notes On Transcription: The plasmid you will transcribe has been linearized by cutting just outside the b-actin insert with Eco RI (see diagram below). This step is important because otherwise the T7 RNA polymerase would transcribe through the b-actin gene and continue around the plasmid transcribing vector sequences. This would not only reduce the amount of label available for labeling b-actin, but might lead to increased background on the blots as vector sequences bound to partially complementary sequences in HeLa cell mRNA.

The following diagrams illustrate the way b-actin is inserted into pBluescript relative and what would happen upon digestion with EcoRI. In this diagram, the top strand of the insert has the same orientation and sequence as actin mRNA.

Actin Sense Strand

Since the T7 RNA polymerase promoter is on the bottom strand, the RNA synthesized will be in an antisense orientation. Cary out the trascription reaction for 2 h at 37^oC. It will not be necessary to add DNase to remove the template DNA at the termination of the reaction, because there is usually a 10 fold excess of RNA over template DNA following transcription, so that the small amount of DNA that might potentially renature with the RNA probe can be ignored.

It would not matter which strand you transcribed for doing a Southern, but is critically important when producing probes for Northern Blotting. Why?

You will be provided with a test strip on which you will spot a 1:3 dilution series of your experimental probe to compare to a dilution series of a prespotted control probe we will provide you.

	A	B	C	D	E
Control Strip	300 pg/ml	100 pg/ml	30 pg/ml	10 pg/ml	3 pg/ml

	A	B	C	D	E
Your Ideal Probe Strip	~300 pg/ml	~100 pg/ml	~33 pg/ml	~10 pg/ml	~3 pg/ml

You will follow the instruction in Protocol 5.12, step 1 on how to dilute and spot your probe so that you can estimate how much probe you synthesized. For detection, follow the instructions in the Colorimetric Detection Method steps 1-4 (Note that this is essentially the same process used for assessing the random primed DNA probe).

3) After you have determined the probe concentration, store the probe at -70^oC.

Expression (Continued from Subcloning)

1) Prepare minipreps of two of your recombinants that were cloned in NovaBlue cells using Protocol 4.24. Note: The Resuspension Buffer we are providing you does NOT have RNAse in it. Each quadrant will be provided with a separate tube of DNAse-free Rnase at 10 mg/ml. You should add 6 ml of this solution to your 1.2 ml of Resuspension Buffer before using it to obtain a final concentration of 50 ug/ml RNase. You should obtain about 1-3 m