01/07/09 Lab 1 - Tissue Extraction I



List of Supplies and Equipment

Background - RNA Extraction
You will isolate RNA from whole tissue using **TriReagent**. TriReagent allows for separation of RNA from other cellular components, including DNA. There are three primary components of TriReagent that allow this to happen. The first is guanidine isothiocyanate which is a potent protein denaturant, the second is phenol, and the third is pH.

Guanidine isothiocyanate denatures proteins, such as the highly abundant histones that coat DNA. Even more importantly, RNases are denatured. This denaturing action allows for better access of phenol (an organic solvent) to cellular proteins and improves its ability to keep the proteins insoluble. The pH of TriReagent is acidic. The low pH keeps DNA out of solution while RNA remains soluble.

After homogenizing/lysing your tissue in TriReagent, chloroform (another organic solvent) will be added to your sample to allow for separation of the phenol and insoluble cellular components (DNA, proteins) from soluble cellular components (RNA). This will result in three distinct layers: the organic phase (the bottom portion), the interphase (layer of cell debris) and the aqueous phase (the top portion). The aqueous phase (the RNA) can then be easily isolated.

The RNA can be precipitated and washed to remove residual phenol and salt carryover. Then the RNA can be resuspended in a suitable solution and quantitated.

RNA is quantitated using a spectrophotometer and measuring the absorbance of your RNA sample at 260nm (A260). The concentration of your sample is calculated with the following equation:
[RNA] = 40ug/mL x A260 x Dilution Factor

In addition to the A260, absorbance at 230nm and 280nm should be taken. The ratio of A260 to each of these absorbances can be used to assess the purity of your RNA. Various substances will absorb at 230nm, which will indicate carryover of phenol, ethanol or high salt in your sample. Proteins generally absorb light at 280nm. For clean RNA, A260/A280 should range between 1.8-2.0. The A260/A230 should range between 1.5-2.0 for clean RNA.

IMPORTANT NOTES

1. Wear clean gloves - For your own safety as well as the integrity of your RNA samples, you must wear gloves throughout this week's lab. Phenol and chloroform are nasty, cuastic chemicals, so gloves are necessary when handling anything that comes in contact with either reagent. Additionally, RNases are constantly secreted from your skin and can easily enter, and subsequently degrade, your RNA sample.

2. Phenol/Chloroform Handling and Disposal -
A. Handling - You must wear gloves, safety glasses and lab coats at all times! These chemicals have potential to do damage to clothing and exposed body parts. TriReagent may be used on the benchtop, but be aware that it is caustic, very volatile and has a very strong odor. Chloroform must only be used in a fume hood! It is extremely caustic, volatile, and inhalation of fumes can be dangerous.
B. Disposal - All tips/tubes/gloves that come in contact with phenol/chloroform must be disposed of in the "Solid Phenol/Chloroform Waste" container found in the fume hood. None of this type of waste should be discarded in regular trash! Any liquids that have phenol/chloroform must be disposed of in the "Liquid Phenol/Chloroform Waste" container found in the fume hood. None of this type of waste should be disposed of down the drain or in the regular trash!

3. RNA Handling - Due to the prevalence of RNases, gloves should be worn at all times when handling your samples. Samples should also be stored on ice at all times (to reduce the activity of any contaminating RNases remaining in your sample), unless otherwise noted.

4. Razor Blades Handling and Disposal -
A. Handling - Obviously, these are extremely sharp. Use them with extreme caution. Pay careful attention to what you are cutting. Only cut tissue that is on a flat, stable surface. Do NOT attempt to cut anything with a razor blade while holding the object in your hand!
B. Disposal - Razor blades MUST be disposed of in the available "Sharps" container! The "Sharps" container is bright red and easily visible. If you cannot find it, ask the TA. Under no circumstances are razor blades to be disposed of in the regular trash!


RNA ISOLATION PROTOCOL
1. Add 500uL of TriReagent to a 1.5mL snap cap tube. Store on ice.
2. Using a clean razor blade, cut a piece of frozen tissue weighing between 50-100mg and add to tube containing TriReagent.
Abalone Sample 07-19-01 (1) = 0.058g
Abalone Sample 06:5-28 (2) = 0.047g
 These samples were gill tissue Due to the lack of material, the same forceps were used for all samples, so there may be some cross-contamination.
3. Carefully homogenize the tissue using a disposable pestle.
4. Add an additional 500uL of TriReagent to the tube and close the tube.
5. Vortex vigorously for 15s.

----- Stop here for Lab 1 and freeze sample at -80 -- I'm not sure if this matters, but the samples sat at room temperature for the length of the lab.


Background - Protein Extraction
You will isolate cellular protein from whole tissue using CelLytic MT. This is a proprietary reagent that contains a mixture of salts and detergents to effectively disrupt lipid membranes, lyse cells, and buffer the cellular proteins at the appropriate pH. This solution will also be supplemented with a cocktail of protease inhibitors to minimize the impact of the numerous proteases that are ubiquitous within all cells.
After extraction, you will determine the concentration of proteins in your sample. Due to the high variability of protein structures, these molecules do not uniformly absorb light at any specific wavelength like nucleic acids. We will use the Bradford Assay to determine the concentration of proteins in your sample. This is a colorimetric assay that uses a reagent (Coomassie Blue) that interacts with proteins. When the reagent is mixed with a solution containing proteins, the solution will turn different intensities of blue depending on the amount of proteins present in the sample. This blue dye absorbs at 595nm and the absorbance can be directly correlated to a specific amount of protein present in the sample when compared to a standard curve (already determined for you).

IMPORTANT NOTES

1. Wear clean gloves - Proteases are present on you skin and are detrimental to the integrity of your samples.
2. Mixing - The Bradford assay works best when samples are mixed well. Invert tubes frequently during incubations, and immediately before measuring absorbance to ensure accurate absorbance readings.


PROTEIN EXTRACTION PROTOCOL
1. Add 0.5mL of CelLytic MT solution to a 1.5mL snap cap tube.
2. Add 25mg of your tissue to the tube.
Abalone Sample 06:5-27 (1) = 0.030g Abalone Sample 06:5-27 (2) = 0.030g I used heart tissue for the protein extraction
3. Homogenize the tissue with a disposable pestle.
4. Close the tube and invert the tube several times.
5. Spin the tube in a refrigerated microfuge for 10mins. @ max speed.
6. While spinning, label a fresh tube with the word "Protein", source organism/tissue, your initials, and today's date.
7. Carefully transfer supernatant to labeled tube and store tube on ice.
8. To a fresh tube, add 1.5mL of Bradford reagent.
9. To this same tube, add 30uL of your protein extract.
10. Invert the tube several times and then incubate at RT for 10mins.
The samples were removed from the microfuge at 3:10pm and mine were measured at 3:45pm, incubating over 3x longer than the necessary 10 minutes.
11. Mix the tube several times and transfer 1mL to a plastic, disposable cuvette. To begin, we zeroed the spectrometer using a 1mL sample of blank solution. Blank solution: in 1.5mL tube, add 1.5mL Bradford reagent and 30uL of CelLytic MT solution.
12. Measure the absorbance at 595nm and record the value.
13. Remove the cuvette from the spectrophotometer. Using a P1000 set to 1mL, carefully pipette the solution in the cuvette up and down a couple of times to mix.
14. Measure the absorbance at 595nm and record the value.
15. Repeat steps 13 and 14.
16. Average the three absorbance values you recorded.
Sample 1: 1.396, 1.385, 1.383, average = 1.388 Sample 2: 1.204, 1.212, 1.209, average = 1.208
17. Plug your average absorbance that you just calculated into the following equation to determine the concentration (ug/mL) of protein in your sample:

[sample] = (Avg. OD595 - 0.04 + 0.0403)/0.00007
[Sample 1] = (1.388 - 0.04 + 0.0403)/0.00007 = 19382.8571 [Sample 2] = (1.208 - 0.04 + 0.0403)/0.00007 = 17261.4286 I'm not sure if I did that right, because those numbers are huge! But I followed the posted equation...
18. Write the concentration on your tube and place tube in TA's ice bucket. Your sample will be stored @ -80C.


Background - Primer Design
Primers, or oligonucleotides (oligos), are short stretches of DNA that are used to direct DNA polymerases to amplify specific regions of larger DNA molecules. Oligos are synthesized by various manufacturer's to contain the precise sequence requested by the customer.

Here is a list of things to take into consideration when designing primers. Although none of these are absolute, they will help ensure your primers will hybridize to your target sequence with the best efficiency.

1. 18-30 bases in length.
2. Melting temp. (Tm) of primers should be within 2C of each other.
3. Avoid primer dimers and primer hairpins
4. Avoid high G/C stretches, particularly at the 3' end
5. G/C clamp at 3' end of primers.

Primer design is most commonly done via computerized means and the algorithms used take the above rules into consideration. Of course, the user always has the opportunity to adjust the parameters that define how primers are designed by the software. There is a great deal of software available for primer design. Integrated DNA Technologies (IDT), a manufacturer of oligos, provides an excellent, free suite of oligo design and anlysis tools. It is a web-based software called SciTools and can be found at their website: www.idtdna.com

The software will allow you to enter a full DNA sequence and then define what region(s) you would like to amplify, the ideal size of the amplicon (PCR product), the ideal length of the oligos, etc. Additionally, after you have selected some proposed primers, you can compare melting temps, G/C content, primer dimer/hairpin probabilitites, etc.

For my RNA sample, I'm using abalone gills, and would like to compare and focus on immune response.

1/14/09 Lab 2 - Tissue Extraction II

*


List of Supplies and Equipment:

RNA ISOLATION PROTOCOL (see also Lab 1 **Tissue Extraction I** and **Manufacturer Protocol**)

1. Turn on heating block to 55C. Also turn on spectrophotometer.
2. Add 500uL of TriReagent to a 1.5mL snap cap tube. Store on ice.
3. Cut a piece of frozen tissue weighing between 50-100mg and add to tube containing TriReagent.
4. Carefully homogenize the tissue using a disposable pestle.
5. Add an additional 500uL of TriReagent to the tube and close the tube.
6. Vortex vigorously for 15s.

----- Stop here for Lab 1 and freeze sample at -80
Turn on heating block to 55C.

7. Incubate tube at room temperature (RT) for 5 mins.
8. In the fume hood, add 200uL of chloroform to your sample and close the tube. NOTE: Due to the high volatility of chloroform, pipetting needs to be done carefully and quickly. Have your tube open and close to the container of chloroform before drawing and chloroform into your pipette tip.
9. Vortex vigorously for 30s. You are vortexing correctly if the solution becomes a milky emulsion.
10. Incubate tube at RT for 5 mins.
11. Spin tube in refrigerated microfuge for 15 mins. @ max speed.
12. Gently remove tube from microfuge. Be sure not to disturb the tube.
13. Slowly and carefully transfer most of the aqueous phase (the top, clear portion) to a fresh microfuge tube. Do NOT transfer ANY of the interphase (the white, cell debris between the aqueous and organic phase).
14. Close the tube containing the organic and interphase and properly dispose of the liquid inside the tube as well as the tube itself at the end of the lab.
15. Add 500uL isopropanol to the new tube containing your RNA and close the tube.
16. Mix by inverting the tube numerous times until the solution appears uniform. Pay particular attention to the appearance of the solution along the edge of the tube. If mixed properly, it should no longer appear viscous/"lumpy".
17. Incubate at RT for 10 mins.
18. Spin in refrigerated microfuge at max speed for 8 mins.
19. A small, white pellet (RNA and salts) should be present. If not, do not fret. Continue with procedure.
20. Remove supernatant.
21. Add 1mL of 75% EtOH to pellet. Close tube and vortex briefly to dislodge pellet from the side of the tube. If the pellet does not become dislodged, that is OK.
22. Spin in refrigerated microfuge at 7500g for 5mins.
23. Carefully remove supernatant. Pellet may be very loose. Make sure not to remove pellet!
24. Briefly spin tube (~15s) to pool residual EtOH.
25. Using a small bore pipette tip (P20 or P200 tips), remove remaining EtOH.
26. Leave tube open and allow pellet to dry at RT for no more than 5mins.
27. Resuspend pellet in 100uL of 0.1%DEPC-H2O by pipetting up and down until pellet is dissolved.
28. Incubated tube at 55C for 5mins. to help solubilize RNA.
29. Remove tube from heat, flick a few times to mix and place sample on ice. This will be your stock RNA sample.
30. Quantitate RNA yield using spectrophotometer.

RNA QUANTIFICATION

1. Obtain two disposable plastic cuvettes: one for a blank and another for your RNA sample.
2. Label both cuvettes at the very TOP of the cuvettes.
3. Add 1mL 0.1%DEPC-H2O to a fresh 1.5mL snap cap tube. This will be your blank.
4. To a fresh 1.5mL snap cap tube, add 990uL 0.1%DEPC-H2O and 10uL of your RNA sample. Mix well by inverting tube multiple times.
5. Transfer these two samples to their respective cuvettes.
6. Ensure wavelength of spectrophotometer is set to 260nm.
7. Insert blank cuvette as demonstrated and zero this sample.
8. Remove blank cuvette from spectrophotometer and insert your RNA sample cuvette.
9. Record the A260 value displayed.
10. Adjust the wavelength and record the values at 230nm and 280nm.
11. Calculate RNA concentration (conversion: 1 A260 unit = 40 ng/uL single-stranded RNA), total RNA yield, A260:A280, and A260:A230.
12. Clearly label your stock RNA sample with the word "RNA", source organism/tissue, your initials, today's date and the concentration in ug/uL.
13. Place tube in ice bucket at the front of the lab. Store your samples at -80C.

Background - SDS - Polyacrylamide Gel Electorophoresis (SDS-PAGE)

SDS-PAGE is the process of separating proteins from one another on the basis of molecular weight. A mixture of proteins is subjected to an electric field and pulled through a polyacrylamide matrix towards the cathode. However, proteins must be treated prior to separating them in this manner. This is because the charge on any given protein is dependent upon the amino acid sequence as well as the pH of the solution. Thus, a crude protein extract from cells or tissue contains a hetergenous mixture of proteins with varying charges. Without treating them in some manner, the proteins will migrate independent of their molecular weight. Additionally, proteins have various tertiary or quarternary structure that can influence the rate at which they migrate in an electric field. In order to address these issues, protein samples are prepared in a specific fashion to linearize the proteins and impart the same charge to all proteins in the sample to ensure that they become homogenous and their migration rate during SDS-PAGE is solely due to molecular weight.

Protein samples are combined with a reducing sample buffer. This reducing sample buffer contains sodium dodecyl sulfate (SDS), B-mercaptoethanol, glycerol, Bromophenol blue and a buffer. SDS imparts an overall negative charge to all the proteins in a sample. This ensures that all of the proteins will migrate in the same direction (towards the cathode) when placed in an electric field. B-mercaptoethanol is a reducing agent. It accepts electrons from disulfide bonds formed between two cysteine residues. It serves as one step to help break any tertiary or quartenary structure of proteins in the sample. Gycerol simply serves as a sinking agent for your sample. It has a greater density and will allow your sample to sink into the buffer contained in the wells of the gel. Bromophenol blue is a negatively charged dye that allows one to visually track the migration of your samples through the polyacrylamide gel. Bromophenol blue migrates at the same rate of proteins ~5-7kDa. Finally, the buffer is present to maintain the appropriate pH for your sample. Once samples have the appropriate amount of reducing sample buffer, they are boiled. Boiling causes the proteins to fully denature, eliminating any tertiary or quartenary structure and leads to linear chains of amino acids.

Samples are then run through polyacrylamide gels. Traditionally, a single polyacrylamide gel is actually comprised of two gels with different percentages of polyacrylamide, pH and buffer. The top portion of the gel, relative to the bottom portion of the gel, has a lower percentage of acrylamide, a lower pH and a lower concentration of buffer. This gel is referred to as the stacking gel. The bottom gel has higher mounts of all three components listed above and is called the running gel.

The low percentage of polyacrylamide in stacking gels allows all the proteins in the sample, regardless of molecular weight, to quickly and easiy migrate through the gel. When the samples begin to enter the lower gel containing a higher percentage of polyacrylamide (running gel), the proteins are now all "stacked" upon one another. This allows all of the proteins in a sample to enter the running gel at essentially the same time. Additionally, the differences in pH and buffer content between the stacking and running gels leads to a local increase in voltage around the sample, which helps drive the sample from solution in the well into the polyacrylamide matrix of the stacking gel.

After SDS-PAGE is complete (when the dye front has reached the bottom of the gel), the gel is either set up for Western blotting (see Lab #4) or is stained to reveal the proteins. There are a number of stains that can be used, depending on the sensitivity needed to visualize proteins of interest, but we will use Coomassie Brilliant Blue. This is a non-selective stain, meaning it binds all proteins regardless of their amino acid makeup. Additionally, it is cheap and rather sensitive. The dye will initially turn the entire gel blue and even a short exposure (5 minutes) often results in over staining. Thus, it is necessary to destain the gel to wash the dye out of the areas of the gel where no protein is present. After destaining, the proteins should appear as blue/purple bands and the rest of the gel should remain relatively clear.

PROTEIN GEL PROTOCOL - See also **Manufacturers Protocol / Manual: Precise™ Protein Gels**

1. Begin boiling water on hot plate.
2. Thaw you protein extract from last week. Mix well by inverting tube several times.
3. In a fresh, 1.5mL SCREW CAP tube add 15uL of your protein sample and 15uL of 2X Reducing Sample Buffer.
4. Mix sample by flicking. Briefly centrifuge (10s) to pool liquid in bottom of tube.
5. Boil sample for 5 mins.
6. While sample is boiling, observe assembly of gel box and gels. Rinse gel wells thoroughly as demonstrated.
7. When sample is finished boiling, immediately centrifuge for 1min. to pool liquid.
8. Slowly load your entire sample into the appropriate well using a gel loading tip.
9. Put lid on gel box and plug electrodes into appropriate receptacles on the power supply.
10. Turn power supply on and set voltage to 150V. Run for 45mins.
11. Add ~150mL (does not have to be measured - just need enough to cover the gel) of Coomassie Stain to a designated container.
11. Turn off power supply and disconnect gel box from power supply.
12. Remove lid from gel box.
13. Disengage the tension wedge.
14. Remove gel from gel box.
15. Carefully crack open cassette to expose gel.
16. Trim wells at top of gel.
17. Notch a designated corner of the gel to help you remember the correct orientation of the gel (i.e. which is the top/bottom of the gel, which is the right/left side(s) of the gel)
18. Place gel into container with Coomassie Stain.
19. Incubate on shaker/rocker for 5 mins.
20. Carefully pour stain back into original container. Be careful not to dump out gel!
21. Rinse gel briefly with 10% acetic acid and pour this wash down the drain.
22. Add ~250mL (no need to measure) 10% acetic acid to container with gel. Incubate on shaker/rockers for 15mins. Change out buffer and repeat until bands become clearly visible. This may need to incubate O/N. If so, cover container with plastic wrap and leave on shaker/rocker.



2/4/08 Lab 3 Reverse Transcription and PCR, and Lab 5 Quantitative PCR

Lab 3 Reverse Transcription and PCR

Quantify RNA
Reverse Transcribe RNA to complementary DNA
Perform PCR

List of Supplies and Equipment:
PCR tubes (0.5 ml; thin walled)
1.5 ml microfuge tubes (RNAse free)
RNA samples (student provided)
AMV RT
Nuclease Free water
filter tips
thermal cycler
kim wipes
2x Immomix Master Mix
microfuge tube racks
ice buckets
dNTPs
oligo dT
2x GoTaq Green Master Mix
timers
agarose
Ethidium Bromide (SR Provided)
gel box
camera
DNA ladder

RNA QUANTIFICATION Through measurement on a spectrophotometer
- nuclotides, RNA, DNA (ss and ds) absorb wavelengths at 260 nm and therefore require purification prior to quantification - the ratio of absorbance of 260 nm and 280 nm assesses teh purity of DNA (pure at ~1.8) and RNA (pure at ~2.0) if lower, it indicates teh presence of phenol, protein, or other contaminants that absorb at 280 nm
REVERSE TRANSCRIPTION PROTOCOL
RT-PCR, a lab technique for amplifying a defined piece of RNA using enzyme reverse transcriptase. RT-PCR is used to determine the amount and type of gene expression, and since RNA can't be completed by PCR, the information needs to be reverse transcribed back into it's complimentary DNA (cDNA). 1. reverse transcribes to cDNA 2. amplification of DNA thru PCR this is different that Q-PCR (real-time PCR)RT-PCR is largely used to diagnose genetic disease and determine the abundance of specific RNA molecules within a cell or tissue to measure gene expression
1. Mix your stock RNA sample by inverting tube several times.
2. Transfer 25ug of your RNA (.25ug of mRNA) to a fresh PCR tube. Bring the volume up to 5uL with PCR water. If necessary, spin tube briefly to pool liquid.
3. Incubate tube at 75C for 5mins in thermal cycler.
4. Transfer tube IMMEDIATELY to ice and incubate for at least 5mins.
5. Make Master Mix

PER RXN
4 ul 5x Buffer (AMV RT Buffer)
8 ul dNTPs (10 mM total) nucleotides
 1 ul __**AMV RTranscriptase**__ This is for use in standard first strand cDNA synthesis reactions. It catalyzes the polymerization of DNA using template DNA, RNA, or RNA-DNA hybrids. It requires a primer
 1 ul Oligo dT Primer - this is a strand of thymine nucleotides that transcribes to the string of adenosine nucleotides at the beginning of a sequence
 1 ul RNase free water
Total = 15 ul

Add MM to tube with diluted RNA in it (total volume now 20 ul)
Vortex
Spot spin
Incubate at RT for 10 min
Incubate at 37C for 1 hr in thermocycler
Heat inactivate @ 95C for 3 min
Spot spin
Leave on ice or store at –20C




PCR
Polymerase Chain Reaction involved amplifying a DNA (genomic or complementary) target using a polymerase, primers (short oligonucleotide), and dNTPs (A, C, T, Gs). In general the reaction is placed in a machine (thermocycler) where a series of temperature changes are performed [Denature (~94), Anneal (primer specific ~50-60), and Extention (~72)].
For this lab you will be using **Promega's GoTaq** Product. Please Read! Promega's GoTaq: ready to use solution with bacterially derived TaqDNA polymerases, dNTPs, MgCl2, and reaction buffers at optimal concentrations for efficient amplification of DNA templates by PCR. For amplification that will be visualized by agarose gel electrophoresis and ethidium bromide staining.

Run each template in duplicate AND make sure to include at least 2 negative controls for each primer (no template).

For a 50μl reaction volume:

Component

Volume

Final Conc.

GoTaq®Green Master Mix, 2X

25

1x

upstream primer, 10μM

0.5–5.0μl

0.1–1.0μM

downstream primer, 10μM

0.5–5.0μl

0.1–1.0μM

DNA template

1–5μl

<250ng

Load reactions into thermocycler.

Make Agarose Gels - This is done in the same method as in the protein gel protocol. I skipped this step and went straight into QPCR to catch up with the rest of the class.

Next Day:
Run out PCR products on agarose gels and photograph.


Warning UV light used to visualize DNA is hazardous.

Lab 5 Quantitative PCR
Perform QPCR

List of Supplies and Equipment:
PCR Plates (white); optically clear caps
1.5 ml microfuge tubes (RNAse free)
Nuclease Free water
filter tips
Opticonthermal cycler
kim wipes
2x Immomix Master Mix
SYBR green dye (or similar dye)
microfuge tube racks
ice buckets
timers
cDNA samples (student provided)

PCR
Polymerase Chain Reaction involved amplifying a DNA (genomic or complementary) target using a polymerase, primers (short oligonucleotide), and dNTPs (A, C, T, Gs). In general the reaction is placed in a machine (thermocycler) where a series of temperature changes are performed [Denature (~94), Anneal (primer specific ~50-60), and Extention (~72)].

Run each template in duplicate AND make sure to include at least 2 negative controls for each primer (no template).

1. Prepare master mix: Prepare enough reactions to run each template in duplicate AND make sure to include at least 2 negative controls for each primer (no template). Add 1 additional reaction to the total to ensure sufficient volume recovery.

For a 50μl reaction volume:

Component

Volume

Final Conc.

Master Mix, 2X (Immomix)

25µL

1x

Syto-13 dye (50uM)

2-5µL

2 - 5µM

upstream primer, 10μM

0.5–5.0μl

0.1–1.0μM

downstream primer, 10μM

0.5–5.0μl

0.1–1.0μM

Ultra Pure Water

to 48uL

NA


18uL x 6 = 108 uLfor the primers, add nMol x 10 = 100 mM, then split into 10mM tubes
 2. Add mastermix to wells of a white PCR plate
3. Thaw cDNA samples.
4. Add 2uL cDNA template to each reaction.
5. Add 2uL of ultra pure water to the negative control wells.
6. Cap the wells securely.
7. If necessary, spin the strips to collect volume in the bottom of the wells.
8. and ensure the lids are clean (wipe with a Kim Wipe) before going into the Opticon.
9. Load the plate, verify the PCR conditions and start the run.

B| 1p, 2p, -C, 1p, 2p, -C -->empty