Evaluating Duplicate Gene Expression
using RT-PCR/SSCP Analysis

Wendel Lab
Department of Ecology, Evolution and Organismal Biology
Iowa State University, Ames, IA 50011, USA

No need for great detail here, but there are a few things to consider. First and foremost, success in the RT-PCR procedure requires relatively clean RNA that is free of genomic DNA; if you don’t have a favorite method for DNA removal, consider the Ambion DNA-FREE reagent, which is easy to use and doesn’t require subsequent heating or phenol/chloroform clean-up (both of which result in a loss of RNA).
Our RT-PCR/SSCP approach was designed around a two-step RT-PCR reaction, where cDNA synthesis and PCR are performed in separate steps. In our lab, we use Ambion’s RETROSCRIPT kit for cDNA synthesis, and we typically include between 1 to 5 ug of total RNA in each cDNA synthesis. The subsequent PCR reaction follows the protocol you have predetermined to work best for the locus using genomic DNA as a template. We use Taq polymerase and buffers from Gibco-BRL (any thermostable polymerase should work), and typically include 1 ul of the uncleaned RT product directly into the PCR step. Subsequent PCR amplification will hopefully give rise to a single band (or a maximum of two) if you are interested in evaluating expression of a single gene; if multiple bands arise from RT-PCR, you may need to first determine whether the presence of paralogues complicates the determination of band homology. Next, the amplification should work well, with both primers matching the target site, because you need abundant gel-purified product (you will gel-isolate the RT-PCR products later, so be prepared for the subsequent sample loss). Finally, primers used to amplify SSCP products should not contain any degenerate sites, as they can give rise to SSCPs.STEP

Again, no need for great detail. After PCR amplification, primers must be removed before SSCP; in addition, the desired band will need to be isolated from additional contaminating bands (if present). Gel isolation of PCR product from agarose gels is the most effective way to desalt and clean DNA, but it is relatively time consuming and expensive. Use the method you like best (We like QIAGEN columns for products < 800 bp, GENE CLEAN for products > 800 bp), but keep in mind that you will only need around 2 - 4 ng of clean, double-stranded PCR product for each SSCP labeling reaction.

The goal of this step is to radioactively label one of the two strands of the original PCR product so that conformation shifts can be readily identified on non-denaturing acrylamide gels. Note that for SSCP reactions, either strand - or both - can be labeled. In fact, it is frequently reported that one strand (say, the strand off the “F” primer) will give rise to confirmation differences while the other strand (“R” in this case) will not reveal polymorphism. There is no way to predict ahead of time which strand will give the best conformation shift, so you should consider labeling both strands, separately and together, when doing the initial screens. As a final note, while this procedure is written for [33P] labeling, we’ve obtained good results using [32P] as well, so long as the specific activity is relatively low (~1000 Ci/mmol). Finally, it may also be possible to evaluate SSCP products using non-radioactive methods, such as silver staining or SyBR green staining, but we haven’t investigated these approaches.

(A) SSCP Template and Primer Mixture:

(B) SSCP Labeling Mixture: Add the following amounts of each reagent per sample –

*Note: Increase this amount if the isotope is > 1 half-life old; you can also use [32P]dCTP but use only half the amount of isotope, or use the same amount of a lower specific activity)

(C) Add 3.5 ul of the SSCP labeling mixture to each template/primer tube. Mix well, add mineral oil if necessary, then run the following PCR cycle:

This section details how to analyze radioactively-labeled SSCP products using manual sequencing gels. Gels can be run at room temperature or at 4 C. Some genes resolve better at one temperature than the other. Urea concentrations range from 2% to 10% - different amounts of urea can give better or worse band separation.

(A) Mixing the Acrylamide Solution – all volumes correspond to 75 mL sequencing gels

Materials needed:
Warm water
10X gel buffer, either TTE or TBE
100 mL Nalgene beaker with stir bar
10% ammonium persulfate, fresh
Urea solid
Acrylamide (FMC-BioWhittaker “MDE” acrylamide solutions)

  1. Microwave water until warm to the touch. This will speed dissolution of urea, if included in the gel buffer (refer to Yip et al.[1999], Biotechniques 27:20-24 for reasons to include denaturants like urea, glycerol or DMSO to the gel) Place a 100 ml Nalgene beaker on the top loading balance; tare the beaker, then add desired amount of urea, Add 18.75 mL of FMC “MDE” polyacrylamide gel solution (or “Long-Ranger”) and 4.75 mL of 10X gel buffer. The gel buffer can either be standard TBE (89 mM Tris/89 mM Boric Acid/2.5 mM EDTA, pH 8.3), or a glycerol-tolerant buffer such as Tris-Taurine-EDTA (“TTE”, Amersham # US71949).
  2. Dilute mixture to 75 mL final volume using warm water. Add a stir bar, and stir until dissolved.

(B) Pouring Gels

  1. Once the gel plates are set up, add 25 ul of TEMED and 250 ul of 10% ammonium persulfate to the acrylamide/buffer solution. Continue stirring for ~ 30 seconds, then remove the beaker from the stir plate.
  2. Carefully tilt the gel away from you and pour the acrylamide solution into the glass plates.
  3. Note: Pour slowly, and down lower edge of the gel. If air bubbles form, stop pouring, then tilt the gel so that the liquid moves below the bubble. Re-tilt the gel so that liquid moves above the bubble (this usually causes the bubble to pop). Repeat if necessary.
  4. Once the gel solution fills the glass plates, slide in the comb with the flat side facing the surface of the gel solution (sharks’ teeth facing up). Push the flat side of the comb into the space only ~ 1cm, and no farther. Clamp the glass plate along the length of the comb using 4 clamps. (if using the small vertical rig, simply place the 20-tooth comb into the acrylamide solution, teeth pointing down).
  5. Examine the gel for leaks. If the gel develops a fast leak (more than 1 drop every 15 seconds), place the gel onto four pipette tip racks so that the glass plates lay horizontally.
  6. Let the solution polymerize at least 1 hour before use. If gel is going to sit overnight, wrap the upper surface of the gel by the comb with plastic wrap to reduce evaporation.

(C) Running Gels

Materials needed:
1L of 1X running buffer per gel, chilled to 4C
Heating Block warmed to 94C
Dry Ice (or ice)
Denaturation/Loading buffer
(95% formamide/20 mM EDTA/ 10 mM NaOH/0.05% Bromphenol blue/0.05% xylene cyanol)
Plastic wrap

  1. Before you begin, prepare 1 liter of 1X gel buffer (either TBE or TTE) for each gel. Put it in the walk-in cold room to equilibrate to 4C. Remove clamps, tape and plastic wrap from the gel. Rinse plates briefly under dH2O to remove excess urea crust. Put all electrophoresis materials into the walk-in cold room. Set plates into the gel rig, with long plate facing out. Tighten down clamps, then add ~ 450 mL of 1X running buffer to both the lower and upper chambers. Gently remove sharkstooth (or 20-T) comb from gel. Denature the SSCP samples by mixing 2 ul of PCR product with 8 ?l of denaturation solution. Heat to 95C for 5 minutes, then snap cool on ice. Store on ice until samples are loaded. Prior to loading samples, rinse the upper surface of the gel with 1X buffer, and then put the sharkstooth comb into place. Push the comb down until the teeth just enter the surface of the gel. Using a sharpie, label each lane on the glass prior to loading samples! Load up to 4 ul of each sample into each gel, and load as rapidly as possible. If lanes appear to be leaking, apply power to the gel and run the samples into the acrylamide. Once loaded, immediately hook up the leads (top = black; bottom = red) and run the gel using between 4 and 7W Constant Power (not constant voltage!). Using 1X TTE running buffer, a setting of 4 W will give a voltage of ~ 500 V and an amperage of ~15 mA. The final run time has to be determined experimentally. Initially, consider running two gels, side by side, with samples loaded in duplicate. Run one gel for 14 hours, and the second gel for 20 hours, and then estimate the “optimal” run time for your gene of interest. When run is complete, shut off power and drain the upper tank. Remove the gel from the rig and lay the sandwich down with the short gel facing up. Using a spatula, gently pry apart the glass plates, removing the gel from the sandwich.
  2. To remove the gel from the glass plate, gently lay a piece of Whatman 3MM paper onto the surface of the acrylamide gel (note – once the gel touches the paper they can’t be separated again, so be sure that you have the paper aligned properly with the gel!). Gently lift the Whatman paper, and the gel will stick to the paper. Dry the gel on the gel dryer for 1 hour at 72C (for best results, use dry ice in the vapor trap).

Analysis of bands can be done either qualitatively to simply verify “presence/absence” of alloallelic transcripts, or semi-quantitatively to estimate the relative abundance of alloallelic transcripts. To accomplish the former, dried gels need only to be exposed to autoradiography film or phosphorimaging screens to reveal the banding patterns (examples shown below). When alloalleles are resolved by SSCP, a minimum of two bands can be observed; if only one band is resolved, there is either silencing at one locus, or the SSCP electrophoresis methods have not resolved the alloalleles. For this reason, it is often helpful to include a positive “genomic DNA” control from the allotetraploid that contains both homoeologous products. For this control to work, the PCR product must be the exact same size as the RT-PCR product (intronless). Presence of two bands in the gDNA control and one band in the cDNA sample provides the most convincing evidence of silencing.

Control experiments show that the phosphorimaging signal derived from homoeologous bands in the allotetraploids corresponds directly to the relative abundance of each transcript. Because of this interesting property, it is possible to accurately quantitate the relative ratio of the two homoeologous transcripts across a range of 1:1 to approximately 100:1. To accomplish this, one only needs to use the quantitation software included with the phosphorimager to measure band “volume” (pixel density). Assuming no PCR bias between homoeologues, the ratio of the signal between the bands (measured as volume units) directly corresponds to the relative ratio of the two transcripts.

Background Information


Denaturation/Loading buffer - 95% formamide/20 mM EDTA/10 mM NaOH/0.05% Bromphenol blue/0.05% xylene cyanol.Per 1.0 ml, mix together the following reagents:

Example SSCP profiles from allotetraploid cotton: Quantitating the ratio of duplicate loci present in a single RT-PCR sample.

If the primers used in the original RT-PCR amplification show no bias with regard to template (that is, they bind to all homoeologues equally well), the ratio of double-stranded PCR product for all homoeologues should be approximately equal to the initial template ratio, which in this case is the ratio of homoeologues in the cDNA pool. To test whether SSCP gels can shed light on the relative ratio of two transcripts in the same cDNA pool, I mixed RT-PCR products from fiber-specific locus “G3” that had initially been amplified from A- and D-genome diploid cottons. This test spanned a wide range of dilution ratios, from 1 part D to 100 parts A, up to 1D:1A, and back to 100D:1A. I chose this locus because SSCP analysis of RT-PCR products revealed a clear polymorphism between these diploid transcripts (actually, RT-PCR products), with the D-genome product migrating faster than the A-genome product.

The gel below shows two of three independent replicates from this quantitation experiment. To briefly summarize, SSCP band signals (detected by phosphorimaging) show a significant correlation between the ratio of the D to A signal (initially measured as “volume” units). In addition, the 95% confidence intervals estimated for all ratios between 1:100 bracketed the true value, and in most cases did not deviate by more than a few percent. These results show that SSCP can be used to estimate the relative ratio two transcripts amplified from a single RT-PCR reaction.

                         Replicate 1                                                             Replicate 2

Dilution Ratios (from L to R)

Fraction, D/A Mean Phosphor Signal, D/A + S.D. 95% Confidence Interval
0D:100A 0.0000 -0.0426 + 0.0424 -0.0906 - 0.0054
1D:100A 0.0099 -0.0115 + 0.0112 -0.0250 - 0.0021
1D:20A 0.0476 0.0458 + 0.0182 0.0252 - 0.0664
1D:7A 0.1250 0.0907 + 0.0169 0.0715 - 0.1098
1D:3A 0.2500 0.2996 + 0.0066 0.2921 - 0.3071
1D:1A 0.5000 0.5248 + 0.0157 0.5070 - 0.5427
3D:1A 0.7500 0.7397 + 0.0223 0.7366 - 0.7427
7D:1A 0.8750 0.8375 + 0.0296 0.7964 - 0.8786
20D:1A 0.9524 0.9183 + 0.0698� 0.8216 - 1.0150
100D:1A 0.9901 0.9922 + 0.0264 0.9557 - 1.0288
100D:0A 1.0000 1.0351 + 0.0364 0.9939 - 1.0763

r2 = 0.99470, m = 1.0109

Protocol updated April 2003. Many thanks to Dr. Keith Adams and Dr. Rich Cronn.