Fabio's+Notebook

Fish 541
I have nanodropped all my samples and calculated dilution values. The only problem is that one of my positive controls has a very high concentration, so it turns out that I would have to take 0.04uL of it to get to the desired concentration. I don't know if we have nano pipettes or if there is another way of doing it. I have calculated to make a solution of 20uL.


 * sample number || dna concentr || 260/280 || uL of DNA || water ||
 * 81 || 18.2 || 1.91 || 5.824176 || 14.17582 ||
 * 165 || 23 || 1.75 || 4.608696 || 15.3913 ||
 * 1267 || 10.6 || 1.47 || 10 || 10 ||
 * 1268 || 10 || 1.77 || 10.6 || 9.4 ||
 * 1289 || 18.8 || 1.72 || 5.638298 || 14.3617 ||
 * 1290 || 16.4 || 1.96 || 6.463415 || 13.53659 ||
 * 1464 || 30.7 || 1.88 || 3.452769 || 16.54723 ||
 * 1757 || 7.3 || 1.32 || 14.52055 || 5.479452 ||
 * 66 || 11.1 || 1.45 || 9.54955 || 10.45045 ||
 * 250 || 6.7 || 2.04 || 15.8209 || 4.179104 ||
 * 844 || 15.9 || 1.49 || 6.666667 || 13.33333 ||
 * 860 || 5.3 || 1.72 || 20 || 0 ||
 * 889 || 37.5 || 1.81 || 2.826667 || 17.17333 ||
 * 1001 || 7.1 || 1.58 || 14.92958 || 5.070423 ||
 * 1296 || 23.9 || 1.92 || 4.435146 || 15.56485 ||
 * 1481 || 16.8 || 1.85 || 6.309524 || 13.69048 ||
 * lactobacillus positive control || 58.7 || 2.02 || 1.805792 || 18.19421 ||
 * clostridium positive control || 2702.4 || 1.92 || 0.039224 || 19.96078 ||
 * lactobacillus positive control || 58.7 || 2.02 || 1.805792 || 18.19421 ||
 * clostridium positive control || 2702.4 || 1.92 || 0.039224 || 19.96078 ||
 * clostridium positive control || 2702.4 || 1.92 || 0.039224 || 19.96078 ||

UPDATE. From my original proposal I have abandoned the idea of testing for pathogens and I will only be testing for commensal/beneficial bacteria. I have picked 16 fecal samples of caribou, all adult males. For each sample I alraedy have cortisol levels. I have selected 8 samples with very low cortisol levels and 8 samples with high cortisol. I have ordered 4 pair of primers: 3 pairs target lactobacillus, bacteroides, and clostridium which are all abundant in mammals guts and feces. In addition I have a primer pair that targets Akkermansia which is a genus made of only one species. This species has been associated with mucosal degradation and bowel inflammation and stress.For the three genera I have positive controls obtained for Dr. Marilyn Roberts from the Environmental and Health Science Department. For each sample I have extracted DNA and I will perform conventional PCR followed by agarose gel. For each sample I have 50μl of DNA extract.

Lab procedure

1. Quantify DNA of fecal samples using the nanodrop. For the positive controls I will have to extract DNA from the Bacteroides. For Clostridium and Lactobacillus I already have DNA extracted and only need to check for DNA concentration.

__** DNA Isolation (DNazol) **__

 * Supplies and Reagents **
 * micropipettes (1-1000 µL)
 * sterile filter pipette tips (1-1000 µL)
 * 1.5 mL microfuge tubes
 * microcentrifuge tube rack
 * microcentrifuge (room temperature)
 * razor blades
 * vortexes
 * DNazol
 * 100% ethanol
 * 75% ethanol
 * 0.1% DEPC water
 * kim wipes
 * Nanodrop


 * Procedure Background **
 * DNazol uses a guanadine-detergent lysing solution to hydrolyze RNA and selectively precipitate DNA from the cell. During the extraction, the cells in the sample are lysed and DNA is precipitated using ethanol. DNA is then solubilized using water or NaOH. The genomic DNA recovery should be between 70 and 100% using this protocol.


 * DNazol is an irritant, so please be careful and wear protective gear when using it.
 * Centrifuging in step 3 removes insoluble tissue fragments, partially hydrolyzed RNA, and excess polysaccharides. This step is necessary to get higher quality DNA.
 * If you have trouble removing your DNA precipitate in step 8, you can centrifuge it at 5,000 x g for 5 minutes (room temperature) so that the DNA will form a pellet.
 * We will "spec" our DNA after extraction. Purified DNA should have a A260/A280 ratio of 1.7-1.9, indicating good quality DNA.


 * DNazol Extraction Protocol (Adapted from MRC manual) **
 * 1) Using a sterile pestle, homogenize your tissue sample in 0.5 mL of DNazol in a 1.5 mL sterile microfuge tube. After the tissue is homogenized, add 0.5 mL more of DNazol and mix well.
 * 2) Let your sample incubate for 5 minutes at room temperature.
 * 3) Spin your sample at 10,000 x g (room temp) for 10 minutes.
 * 4) Transfer your supernatant to a new, labeled tube.
 * 5) Add 0.5 mL of 100 % ethanol to your sample.
 * 6) Mix your sample by inverting your tube 5-8 times.
 * 7) Store your sample at room temperature for 1 minute.
 * 8) Your DNA should form a cloudy precipitate. Remove the DNA and put in a new tube using your pipette.
 * 9) Let your sample sit at room temp for 1 minute and remove the rest of the lysate (liquid that is not DNA).
 * 10) Wash your DNA with 1 mL of 75% ethanol: Pipette the ethanol into your DNA tube, invert 6 times, and let sit for 1 minute. Remove the ethanol from the tube and repeat.
 * 11) If there is ethanol left at the bottom of your tube after the second wash, remove with a small pipette.
 * 12) Add 300 µL of 0.1% DEPC water to your DNA and pipette up and down multiple times to dissolve.
 * 13) Bring your DNA sample up to the Nanodrop to quantify.


 * DNA Quantification **


 * 1) Pipette 2µL of 0.1%DEPC-H20 onto the Nanodrop pedestal and lower the arm.
 * 2) Select "dsDNA" from the pulldown menu
 * 3) Click "Blank", to zero the instrument. NOTE: steps 1 and 2 only need to be done once for the whole class.
 * 4) Pipette 2µL of your DNA sample onto the Nanodrop pedestal and lower the arm
 * 5) Click "Measure". Record your DNA concentration (ng/µL), A260/280 ratio and A260/230 ratio. NOTE: The Nanodrop uses the Beer-Lambert Law to calculate DNA concentration for you.
 * 6) Raise the arm and wipe off you sample with a KimWipe
 * 7) Clearly label your stock DNA sample with the word "DNA", source organism/tissue, your initials, today's date and the concentration in ug/uL.
 * 8) Store sample at -20ºC.

2. Conventional PCR


 * Polymerase chain reaction **


 * Supplies and Equipment: **


 * Micropipettes (1-1000 μl)
 * Sterile filter pipette tips (1-1000 μl)
 * Tip waste jar
 * PCR tubes (0.5 ml; thin walled)
 * 1.5 ml microcentrifuge tubes (RNAse free)
 * cDNA (student provided) - in my case this would be DNA
 * dNTPs
 * 2x GoTaq Green Master Mix
 * Primers
 * Nuclease Free water
 * thermal cycler
 * Kimwipes
 * microfuge tube racks
 * PCR tube racks
 * ice buckets
 * Lab coat
 * Safety glasses
 * Gloves


 * Procedure background **

The polymerase chain reaction involves selective amplification of a DNA (genomic or complementary) target using the enzyme polymerase (after which the method is named), primers (short oligonucleotides), and dNTPs (A, C, T, and G). The method relies on thermal cycling, which consist of repeated heating and cooling of the reaction for DNA melting and enzymatic replication of the DNA. These heating and cooling cycles are comprised of three primary steps. A single cycle begins with denaturation at ~94°C and during this step the DNA is melted or rather unwound and the strands are pulled apart resulting in single stranded DNA. This is followed by an annealing step at ~50-60°C where the primers anneal to the target sequence. After which there is an extension step at ~72°C where nucleotides DNA polymerase synthesizes a new DNA strand by adding dNTPs that are complementary to the template. As PCR progresses, the newly generated DNA is also used as a template for replication, setting in motion a chain reaction in which the DNA template is exponentially amplified. This process allows us to generate thousands to millions of copies of a particular DNA sequence with only a single or a few copies of a piece of DNA.

For this lab we will be using Promega’s Go-Taq green master mix, please read the __ manufacture’s protocol __.


 * PCR PROTOCOL **

You will be preparing 50 µl PCR reactions: count how many total samples you have ( = x), add two negative controls (you will use nuclease free water instead of cDNA template), and add one extra reaction to account for pipetting error. For example, if I have 3 cDNA samples, I will calculate my master mix for 6 reactions.

I have 16 sample plus 1 positive controls, repeated 4 times one for each bacteria. So for each primer pair I should I have 18 reactions. (Should I include 1 or 2 positive controls per bacteria). For the Akkermansia I do not have positive control but I will include two negative controls with just water.

1. Make a reaction mastermix in a 1.5 ml microcentrifuge tube labled "MM" and with your initials.

2.Prepare your mastermix by first calculating the total volume required for each component of the mastermix, then pipette each reagent and mix well. The following volumes are what you would need for __a single__ reaction:


 * Reagent || 1.reaction || x.reactions ||
 * 5x GoTaq Green buffer || 12.5 µL || 237.5 ||
 * 10 µM forward primer || 1 µL || 19 ||
 * 10 µM reverse primer || 1 µL || 19 ||
 * dna template || 2 uL ||  ||
 * nuclease-free water || 8.5uL || 161.5 ||
 * nuclease-free water || 8.5uL || 161.5 ||

2. Pipette 48 ul of your master mix in to each of your 0.5 ml PCR tubes labeled with your sample name and with your initials

3. Add 2 ul of the appropriate template to each tube and mix via pipetting

4. Spin tubes to pool liquid at the bottom of the tubes. Load reactions into thermocycler making sure the caps are tightly secured. 5. Your samples will be put through the following thermal cycling profile and then stored at -20°C afterwards.

I will need to adjust the annealing temperature based on the ones I found in the literature where I have taken the primers.
 * Step || Temperature || Time || Cycles ||
 * Denaturation || 95C || 5 min || 1 ||
 * Denaturation || 95C || 30 sec || 40 ||
 * Annealing || 55C || 30 sec ||^  ||
 * Extension || 72C || 90 sec ||^  ||
 * Final extension || 72C || 3 min || 1 ||
 * Hold || 4C || ∞ || 1 ||

3. Run PCR products in agarose gel

If possible I would like to make a gel comb with at least 10 wells or even better 18. I am just unclear at what phase the PCR product comes into place gel procedure. If there is a 10 wells gel I plan to use 4 samples with high cortisol and 4 with low to possibly make it more evident in the gel any difference in abundance.


 * Making an agarose gel ( **//** This has been done ahead of time **//** ) **


 * Supplies and Equipment: **


 * Micropipettes (1-1000 μl)
 * Sterile filter pipette tips (1-1000 μl)
 * Tip waste jar
 * 1L flask
 * agarose
 * 1X TAE
 * Ethidium bromide
 * Microwave
 * Gel rigs
 * Kimwipes
 * Lab coat
 * Safety glasses
 * gloves


 * AGAROSE GEL POURING PROCEDURE **


 * 1) Weigh 2g of agarose and mix with 150mL 1x TAE in a 1L flask
 * 2) Microwave solution for ~ 3 minutes. Keep an eye on the solution so that it does not boil over. You want the solution to be clear - no precipitate and no bubbles.
 * 3) Cool solution (you should be able to touch the flask for a few seconds), then add 12uL ethidium bromide(EtBr). WARNING: EtBr is a carcinogen be sure to wear gloves and appropriately dispose tip waste.
 * 4) Mix thoroughly by swirling, then pour into gel tray.
 * 5) Add gel combs. Using a clean pipet tip, pop any bubbles that could get in the way of your PCR product.
 * 6) After gel is set, wrap in plastic wrap (label with your initials and date) and place gel in the fridge if not using immediately.


 * Agarose Gel Electrophoresis **


 * Procedure Background **


 * Nucleic acid molecules are separated by applying an electric field to move the negatively charged molecules through an agarose matrix. Shorter molecules move faster and migrate farther than longer ones because shorter molecules migrate more easily through the pores of the gel. This phenomenon is called sieving.
 * The most common dye used to make DNA or RNA bands visible for agarose gel electrophoresis is ethidium bromide usually abbreviated as EtBr. It fluoresces under UV light when intercalated into DNA (or RNA). By running DNA through an EtBr-treated gel and visualizing it with UV light, any band containing more than ~20 ng DNA becomes distinctly visible. EtBr is a known mutagen, however, so safer alternatives are available.
 * A DNA ladder is a solution of DNA molecules of different lengths used in agarose gel electrophoresis. It is applied to an agarose gel as a reference to estimate the size of unknown DNA molecules
 * If amplification was successful you should see one clear band between 150-400bp (in length depending on your gene) and the negative controls will have no band. If there is a band in the negative control than there might be contamination in your reagents and you can not be sure that the intended gene was actually amplified.
 * Often contamination requires carful rePCR in case the contamination occurred during reaction setup. However, if any reagents are contaminated troubleshooting may be required to obtain a clean PCR product.
 * PCR bands outside of the intended size range could indicate unspecific amplification and will require either optimization of the reaction cocktail and thermal cycling parameters or redesigning the primers.


 * ELECTROPHORESIS PROCEDURE **
 * 1) Place gel in gel box and fill with 1x TAE buffer (to fully cover wells)
 * 2) Remove combs from wells
 * 3) Load 7uL 100bp ladder in far left lane
 * 4) Load 20uL of your PCR sample into the gel (retain the remaining vol at -20ºC)
 * 5) Run gel at ~ 100V for ~ 1hr
 * 6) Visualize the gel on the UV transilluminator

independent project week three - November 12, 2013 This past week I accomplished to select and order primers. Next step is to select 16 samples of caribou feces to test for bacteria. Following DNA extraction I would like do qPCR based on a procedure I found on this publication “Development of an extensive set of 16S rDNA-targeted primers for quantification of pathogenic and indigenous bacteria in faecal samples by real-time PCR” [] for which I report here the methodology

Real-time PCR optimization and conditions

Performance, optimal annealing temperatures of the PCR primer pairs and expected product sizes (Table 1) were first determined with gradient PCR (Peltier Thermal Cycler PTC-200; MJ Research, Waltham, MA, USA). The amplification reactions were carried out in a total volume of 50 μl, and the standard reaction mixture consisted of 10 mm Tris-HCl (pH 8·8), 150 mm KCl, 0·1% Triton X-100, 1·5 mm MgCl2, 200 μm each dNTP, 1 μm each primer and 1·2 U Dynazyme II polymerase (Finnzymes, Espoo, Finland). Amplification programme included an initial denaturation step at 95°C for 5 min followed by 30 cycles of denaturation at 95°C for 15 s, primer annealing at 50–70°C for 20 s and primer extension at 72°C for 45 s, with final extension step at 72°C for 5 min. The specificity of the primer pairs was confirmed by employing an extensive set of predominant and pathogenic GI tract bacterial species as a negative control. The PCR products were subjected to electrophoresis on agarose gels and stained with ethidium bromide.

SYBR Green PCR amplifications were performed using an iCycler iQ Real-Time Detection System (Bio-Rad) associated with the iCycler Optical System Interface software (version 2·3; Bio-Rad). All PCR experiments were carried out in triplicate with a reaction volume of 25 μl, using iCycler IQ 96-well optical grade PCR plates (Bio-Rad) covered with iCycler optical-quality sealing film (Bio-Rad).

The efficiency of PCR amplification was optimized for each primer pair, using various MgCl2 concentrations with a dilution series of genomic DNA from the target test species. Optimal reaction conditions for each PCR assay are summarized in Table 1. The reaction mixtures for the optimized SYBR Green I-based assays consisted of a 1 : 75 000 dilution of SYBR Green I (Molecular Probes, Eugene, OR, USA), 10 mm Tris-HCl (pH 8·8), 150 mm KCl, 0·1% Triton X-100, 2–5 mm MgCl2, 100 μm each dNTP, 0·5 μm each primer, 0·6 U Dynazyme II polymerase (Finnzymes) and either 5 μl of template or water (see Table 1 for detailed information on the concentrations of the reaction components). Amplification involved one cycle at 95°C for 5 min for initial denaturation followed by 35 cycles of denaturation at 95°C for 15 s, primer annealing at the optimal temperatures (see Table 1) for 20 s, extension at 72°C for 30 s and an additional incubation step at 80–85°C for 30 s to collect the fluorescent data. An extensive set of representative GI bacterial species was used as a negative control. To determine the specificity of PCR reactions, melt curve analysis was carried out after amplification by slow cooling from 95 to 60°C, with fluorescence collection at 0·3°C intervals and a hold of 10 s at each decrement. Preparation of PCR standards and quantification of target bacterial DNA in pure cultured or faecal samples by real-time PCR

For construction of standard curves, 10-fold dilution series of between 0·1 pg and 10 ng (ca 30–100 to 3·0 × 106–1·0 × 107 target genomes) from target species genomic DNA preparations were applied for PCR. A mixture of DNA isolated from an extensive set of nontarget test bacteria (100 pg each) was used as a negative control. The standard curves of individual real-time PCR assays were used for quantification of the target bacterial DNA from faecal DNA preparations.

In order to evaluate the reliability of the DNA purification method used in this study, E. coli subgroup-specific 16S primers (Malinen et al. 2003) were applied for quantification of E. coli genomes from serial dilutions of overnight cultured E. coli DSM 6897 with real-time PCR. Cell density of the overnight culture was estimated by viable count on Luria agar (Difco) and microscopic cell count with the Petroff Hausser Counting Chamber (Hausser Scientific Company, Horsham, PA, USA). Cell lysis and DNA extraction were performed as described in ‘extraction and purification of DNA from faecal samples and bacterial cultures’.

Additionally, real-time PCR was used for the quantification of pure cultured cells of certain pathogenic bacterial strains introduced into faecal samples. Bacterial densities in the pure cultures were estimated microscopically with Petroff Hausser Counting Chamber (Hausser Scientific Company). Replicate faecal samples were spiked with 6 × 103, 6 × 104, 6 × 105, 6 × 106, 6 × 107, 6 × 108 or 0 pure cultured cells of Camp. jejuni, Cl. difficile or H. pylori and subjected to real-time PCR analysis. Campylobacter genus-specific PCR assay was used for quantification of Camp. jejuni whereas species-specific PCR assays were used for quantification of the other two target bacteria (Table 1). Collection of bacteria from the faeces, cell lysis and DNA extraction were performed as described in ‘extraction and purification of DNA from faecal samples and bacterial cultures’.

Independent project week two – November 5, 2013

I had hoped this week to order the primers needed for the real-time PCR, however I had to consult to experts from the health science department that are working on got microflora and was not able to obtain an answer on till the end of the week. I have reviewed more literature and have found different sets of primers for all the genera of bacteria I need and should be able now to order the primers. I have settled on the Caribou samples because I will be able to know their hormone levels and be able to handpick the samples from different areas and with different hormone levels. If I am able to select samples and receive primers by the end of the week I could start processing early next week. The most challenging part about picking the primers is that there were different studies that used different primers based on the host for example humans or herbivores but testing for the same genus. This made it confusing because in theory the same species or genus could be tested with the same primers regardless of the host. Apparently though for my study this does not matter because I am not trying to detect the origin of the bacteria but rather to detect their presence.

Independent Project week 1- October 29, 2013

This week we began working on our own independent project. For my project I will be investigating the correlation between stress and poor diet and the presence or absence of pathogens in deer and also the influence on gut microflora. We hypothesize that under eight not adequate diet deer might compromise their microflora and also be exposed to pathogens. This week was dedicated to selecting which bacteria should be investigated. After searching different sources and consulting a biologist whose expertise is pathogens in ruminants I decided to only focus on gut microflora because detecting harmful bacteria without any prior knowledge involves too many unknown variables. I am also evaluating whether I should use the caribou samples because I have stress related data of them, and I do not have any direct measures of stress on deer. I also settled on four genus Bacteroides, Clostridium, Lactobacillus, and Akkermansia. Now I need to order the primers and start picking samples to be processed.

Lab 4 - October 22nd 2013

The focus of the lab was to use the protein sample that was prepared in the previous lab and use it in agarose gel electrophoresis for correct amplification and also using the SDS-PAGE procedure followed by staining and western blotting. These procedures allow to first separate proteins based on molecular weight. However proteins need to be pretreated to avoid migration of protein regardless of their weight, because proteins have a mix of proteins could have proteins sharing the same charge. This lab was mainly run as a group lab as we used one SDS-PAGE apparatus filled with gel. I positioned my protein in slot 4 on the second section. We had to be extremely careful because the quantity of the protein are extremely low. After the SDS-PAGE we extracted the gel containing the proteins and place it between a membrane and filter paper. In this step was critical to avoid touching the membrane and also to avoid air bubbles within the membrane and the paper as air bubbles would hamper the flow of electricity. After this we set the membrane on a blocking solution and set it on rotary shaker.

My protein samples showed in both procedures, and from what I understood the results were acceptable.

REFLECTIONS

It was interesting to have a group lab session where everybody seemed to collaborated and everybody did a little task. It was fascinating to see how proteins can be pulled apart even when they might have such minimal weight differences. I felt though that interpreting the results was challenging as it was difficult to look at such tiny differences in the width of the bands.

The lab procedures follow below


 * Lab Objectives **


 * Run extracted total protein from previous labs on SDS-PAGE Gel/Western Blot
 * Run conventional PCR samples on an agarose gel
 * Download qPCR data and discuss analysis


 * Making an agarose gel (//This has been done ahead of time//) **


 * Supplies and Equipment: **


 * Micropipettes (1-1000 μl)
 * Sterile filter pipette tips (1-1000 μl)
 * Tip waste jar
 * 1L flask
 * agarose
 * 1X TAE
 * Ethidium bromide
 * Microwave
 * Gel rigs
 * Kimwipes
 * Lab coat
 * Safety glasses
 * gloves


 * AGAROSE GEL POURING PROCEDURE **


 * 1) Weigh 2g of agarose and mix with 150mL 1x TAE in a 1L flask
 * 2) Microwave solution for ~ 3 minutes. Keep an eye on the solution so that it does not boil over. You want the solution to be clear - no precipitate and no bubbles.
 * 3) Cool solution (you should be able to touch the flask for a few seconds), then add 12uL ethidium bromide(EtBr). WARNING: EtBr is a carcinogen be sure to wear gloves and appropriately dispose tip waste.
 * 4) Mix thoroughly by swirling, then pour into gel tray.
 * 5) Add gel combs. Using a clean pipet tip, pop any bubbles that could get in the way of your PCR product.
 * 6) After gel is set, wrap in plastic wrap (label with your initials and date) and place gel in the fridge if not using immediately.


 * Agarose Gel Electrophoresis **

// Last week you performed a conventional PCR in class using your reverse transcribed cDNA samples as template and primers your designed. This week we will be checking if amplification was successful using electrophoresis. //


 * Procedure Background **


 * Nucleic acid molecules are separated by applying an electric field to move the negatively charged molecules through an agarose matrix. Shorter molecules move faster and migrate farther than longer ones because shorter molecules migrate more easily through the pores of the gel. This phenomenon is called sieving.
 * The most common dye used to make DNA or RNA bands visible for agarose gel electrophoresis is ethidium bromide usually abbreviated as EtBr. It fluoresces under UV light when intercalated into DNA (or RNA). By running DNA through an EtBr-treated gel and visualizing it with UV light, any band containing more than ~20 ng DNA becomes distinctly visible. EtBr is a known mutagen, however, so safer alternatives are available.
 * A DNA ladder is a solution of DNA molecules of different lengths used in agarose gel electrophoresis. It is applied to an agarose gel as a reference to estimate the size of unknown DNA molecules
 * If amplification was successful you should see one clear band between 150-400bp (in length depending on your gene) and the negative controls will have no band. If there is a band in the negative control than there might be contamination in your reagents and you can not be sure that the intended gene was actually amplified.
 * Often contamination requires carful rePCR in case the contamination occurred during reaction setup. However, if any reagents are contaminated troubleshooting may be required to obtain a clean PCR product.
 * PCR bands outside of the intended size range could indicate unspecific amplification and will require either optimization of the reaction cocktail and thermal cycling parameters or redesigning the primers.


 * ELECTROPHORESIS PROCEDURE **


 * 1) Place gel in gel box and fill with 1x TAE buffer (to fully cover wells)
 * 2) Remove combs from wells
 * 3) Load 7uL 100bp ladder in far left lane
 * 4) Load 20uL of your PCR sample into the gel (retain the remaining vol at -20ºC)
 * 5) Run gel at ~ 100V for ~ 1hr
 * 6) Visualize the gel on the UV transilluminator

**Protein Extraction and Analysis Part 2**


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


 * Supplies and Reagents **


 * micropipettes (1-1000 μL)
 * sterile filter pipette tips (1-1000 μL)
 * sterile gel loading tips
 * 1.5 mL screw cap tubes
 * microcentrifuge tube rack
 * lab coats
 * safety glasses
 * gloves
 * lab pen
 * timers
 * heating block with water bath
 * tube "floatie" (8 tube capacity)
 * glass container for boiling water that can accommodate "floatie"
 * protein gel box (SR provided)
 * SDS/PAGE gels
 * gel loading tips
 * trays for staining gels
 * power supply
 * platform rocker/shaker
 * plastic wrap
 * 2X SDS reducing sample buffer
 * protein ladder marker
 * gel running buffer
 * light box
 * digital camera


 * SDS-PAGE PROTOCOL **

Also see [|Manufacturers Protocol / Manual: Precise™ Protein Gels]


 * 1) // Begin boiling water on hot plate. //
 * 2) In a fresh, 1.5mL SCREW CAP tube add 15uL of your protein stock and 15uL of 2X Reducing Sample Buffer. Return your protein stock to the box in the -20C freezer labeled protein samples.
 * 3) Mix sample by flicking. Briefly centrifuge (10s) to pool liquid in bottom of tube.
 * 4) Boil sample for 5 mins.
 * 5) While sample is boiling, observe assembly of gel box and gels. Rinse gel wells thoroughly as demonstrated.
 * 6) When sample is finished boiling, immediately centrifuge for 1min. to pool liquid.
 * 7) Slowly load your entire sample into the appropriate well using a gel loading tip.
 * 8) Put lid on gel box and plug electrodes into appropriate receptacles on the power supply.
 * 9) Turn power supply on and set voltage to 150V. Run for 45mins. **CHECK YOUR AGAROSE GEL RESULTS. MAKE SURE EVERYTHING IS SET UP FOR WESTERN BLOT.**
 * 10) Turn off power supply and disconnect gel box from power supply.
 * 11) Remove lid from gel box.
 * 12) Disengage the tension wedge.
 * 13) Remove gel from gel box.
 * 14) Carefully crack open cassette to expose gel.
 * 15) Trim wells at top of gel.
 * 16) 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)
 * 17) Proceed to Western Blotting protocol.

== ** WesternBreeze Chromogenic Western Blot Immunodetection ** ==

**Supplies and Reagents**


 * Nanopure water
 * gel staining tray
 * Blocking Solution
 * rotary shaker
 * Primary Antibody Solution
 * Antibody Wash
 * Secondary Antibody Solution
 * Chromogenic Substrate
 * timers
 * lab coats
 * safety goggles
 * gloves
 * SDS-PAGE gel
 * Tris-Glycine transfer buffer
 * filter paper
 * nitrocellulose membrane
 * semi-dry transfer station


 * Western Blot Protocol **


 * This is a lengthy protocol with many incubation steps. There will be one gel for the entire class. As a class, you should assign yourselves steps so that everyone can participate and so that we don't waste time. When it is not your turn to attend to the gel, you can do protein and RNA extractions for your project. **


 * 1) Soak the filter paper, membrane and gel in Tris-Glycine Transfer Buffer for 15 minutes.
 * 2) Assemble the blotting sandwich in the semi-dry blotting appartus:
 * 3) Anode (+++)
 * 4) filter paper
 * 5) membrane
 * 6) gel
 * 7) filter paper
 * 8) cathode (---)


 * 1) Transfer the blot for 30 minutes at 20V
 * 2) Remove the gel from the sandwich and rinse off adhering pieces of gel with transfer buffer.
 * 3) Wash membrane 2 times, for 5 minutes each, with 20 mL of pure water.
 * 4) Put the membrane in the plastic box and add 10 mL of Blocking Solution. Cover and incubate overnight on a rotary shaker set at 1 revolution/second.
 * 5) //Your TA will do the rest of the steps. After class tomorrow you can come and see your results.//
 * 6) Decant liquid.
 * 7) Rinse the membrane with 20 mL of water for 5 minutes, then decant. Repeat.
 * 8) Incubate the membrane in 10 mL of Primary Antibody Solution. Decant the solution.
 * 9) Rinse the membrane with 20 mL of Antibody Wash for 5 minutes, then decant. Repeat 3 times.
 * 10) Incubate the membrane in 10 mL of Secondary Antibody Solution for 30 minutes. Decant.
 * 11) Wash the membrane for 5 minutes with 20 mL of Antibody wash, then decant. Repeat 3 times.
 * 12) Rinse the membrane with 20 mL of pure water for 2 minutes, then decant. Repeat twice.
 * 13) Incubate the membrane in 5 mL of Chromogenic Substrate until a purple band appears. This will occur between 1-60 minutes after adding the Chromogenic Substrate.
 * 14) Dry the membrane on a clean piece of filter paper to the open air.

Lab 3 - October 15th 2013

The lab started from the cDNA sample that was prepared in the Lab 2. The purpose of this lab was to prepare the test tube to perform qPCR. We prepared 4 tubes, two of which were test tubes set up with water without the cDNA and the other two contained cDNA. All tubes contained a master mix made with SsoFast EvaGreen supermix and upstread and downstream primer. Different primers were available to amplify different sections. I chose to test for HSP.


 * qPCR PROCEDURE **

// You will run each template (cDNA) in duplicate in addition to two negative controls (no template) - calculate how many reactions this will be! //

1. Prepare master mix: Prepare enough master mix for your number of reactions +1 to ensure sufficient volume recovery.

// For a 25μl reaction volu // me:
 * **Component** || **Volume** || **Final Conc.** ||
 * Master Mix (SsoFast EvaGreen supermix) || 12.5µL || 1x ||
 * upstream primer, 10μM || 0.5μl || 2.5μM ||
 * downstream primer, 10μM || 0.5μl || 2.5μM ||
 * Ultra Pure Water || 10.5uL || NA ||

2. Add mastermix to wells of a white PCR plate 3. Thaw cDNA samples. 4. Add 1uL cDNA template to each reaction. 5. Add 1uL 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. Ensure the lids are clean and place strips on ice. (I like to wipe the lids with a clean kimwipe) 9. Load the plate, verify the PCR conditions and start the run (this will be done by your TA).

PCR conditions: 1. 95°C for 10 minutes 2. 95°C for 15s 3. 55 °C for 15 s 4. 72°C for 15 s (+ plate read) 5. Return to step 2 39 more times 6. 95°C for 10s 7. Melt curve from 65°C to 95°C, at 0.5°C for 5s (+plate read)

RESULTS

Unfortunately I made a mistake and prepared all the test tubes with cDNA and didn't have any test tubes. However the results were positive with amplification within the desirable range of cycle. In addition though the melt curve shows that the cDNA might contain some gDNA, however this is only one possible hypothesis.

CONSIDERATION

In retrospective this lab was interesting because it helped me visualize through the PCR graphs how the process of amplification works and how it's possible to test if the cDNA/gDNA samples were prepared accurately. It was also quite challenging to deal with such small quantities as 1 microL.

Lab 2 – October 8th 2013

In this lab session we continued with the RNA processing that was started during the first lab. After taking the sample out of the freezer we added chloroform and spun it in the centrifuge to obtain three distinctive layers. Only did top layer was to be taken and set in a new you tube with the addition of isopropanol. Once the test tube was spun again in the centrifuge we obtained a pellet which was then extracted and centrifuged with ethanol. Once the ethanol was removed the pellet was let dry and suspended in 100uL of 0.1%DEPC-water and incubated at 55°C for a few minutes. At this point we skipped the RNA quantification and used part of the RNA solution to prepare a primer solution to produce a cDNA sample. The lab material and detailed instructions follow:


 * RNA Extraction Part 2 **


 * Supplies and Reagents **
 * micropipettes (1-1000 μL)
 * sterile filter pipette tips (1-1000 μL)
 * 1.5 mL microcentrifuge tubes
 * microcentrifuge tube rack
 * <span style="font-family: 'Arial','sans-serif'; font-size: 16px;">lab coats
 * <span style="font-family: 'Arial','sans-serif'; font-size: 16px;">safety glasses
 * <span style="font-family: 'Arial','sans-serif'; font-size: 16px;">gloves
 * <span style="font-family: 'Arial','sans-serif'; font-size: 16px;">lab pen
 * <span style="font-family: 'Arial','sans-serif'; font-size: 16px;">timers
 * <span style="font-family: 'Arial','sans-serif'; font-size: 16px;">ice buckets
 * <span style="font-family: 'Arial','sans-serif'; font-size: 16px;">phenol/chloroform waste containers (liquid/solid)
 * <span style="font-family: 'Arial','sans-serif'; font-size: 16px;">vortex
 * <span style="font-family: 'Arial','sans-serif'; font-size: 16px;">hot water bath
 * <span style="font-family: 'Arial','sans-serif'; font-size: 16px;">Nanodrop spectrophotometer
 * <span style="font-family: 'Arial','sans-serif'; font-size: 16px;">chloroform
 * <span style="font-family: 'Arial','sans-serif'; font-size: 16px;">RNase free water
 * <span style="font-family: 'Arial','sans-serif'; font-size: 16px;">chloroform
 * <span style="font-family: 'Arial','sans-serif'; font-size: 16px;">isopropanol
 * <span style="font-family: 'Arial','sans-serif'; font-size: 16px;">75% ethanol
 * <span style="font-family: 'Arial','sans-serif'; font-size: 16px;">0.1% DEPC treated water
 * <span style="font-family: 'Times New Roman','serif';">RNA EXTRACTION PROTOCOL **

<span style="font-family: 'Times New Roman','serif';"> //Continued from Lab 1// <span style="font-family: 'Times New Roman','serif';"> 1. Turn on heating block to 55°C. <span style="font-family: 'Times New Roman','serif';"> 2. Incubate your homogenized tissue sample (from Lab 1) tube at room temperature (RT) for 5 mins. <span style="font-family: 'Times New Roman','serif';"> 3. 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. <span style="font-family: 'Times New Roman','serif';"> 4. Vortex vigorously for 30s. You are vortexing correctly if the solution becomes a milky emulsion. <span style="font-family: 'Times New Roman','serif';"> 5. Incubate tube at RT for 5 mins. <span style="font-family: 'Times New Roman','serif';"> 6. Spin tube in refrigerated microfuge for 15 mins. @ max speed. <span style="font-family: 'Times New Roman','serif';"> 7. Gently remove tube from microfuge. Be sure not to disturb the tube. <span style="font-family: 'Times New Roman','serif';"> 8. 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 dbris between the aqueous and organic phase). <span style="font-family: 'Times New Roman','serif';"> 9. 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. <span style="font-family: 'Times New Roman','serif';"> 10. Add 500uL isopropanol to the new tube containing your RNA and close the tube. <span style="font-family: 'Times New Roman','serif';"> 11. 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". <span style="font-family: 'Times New Roman','serif';"> 12. Incubate at RT for 10 mins. <span style="font-family: 'Times New Roman','serif';"> 13. Spin in refrigerated microfuge at max speed for 8 mins. When placing your tube in the microfuge position the tube hinge pointing up, away from the center of the microfuge. <span style="font-family: 'Times New Roman','serif';"> 14. A small, white pellet (RNA and salts) should be present. If not, do not fret an continue with the procedure. <span style="font-family: 'Times New Roman','serif';"> 15. Remove supernatant. <span style="font-family: 'Times New Roman','serif';"> 16. 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. <span style="font-family: 'Times New Roman','serif';"> 17. Spin in refrigerated microfuge at 7500g for 5mins. <span style="font-family: 'Times New Roman','serif';"> 18. Carefully remove supernatant. Pellet may be very loose. Make sure not to remove pellet! <span style="font-family: 'Times New Roman','serif';"> 19. Briefly spin tube (~15s) to pool residual EtOH. <span style="font-family: 'Times New Roman','serif';"> 20. Using a small pipette tip (P10 or P20 tips), remove remaining EtOH. <span style="font-family: 'Times New Roman','serif';"> 21. Leave tube open and allow pellet to dry at RT for no more than 5mins. <span style="font-family: 'Times New Roman','serif';"> 22. Resuspend pellet in 100uL of 0.1%DEPC-H2O by pipetting up and down until pellet is dissolved. <span style="font-family: 'Times New Roman','serif';"> 23. Incubated tube at 55C for 5mins. to help solubilize RNA. <span style="font-family: 'Times New Roman','serif';"> 24. Remove tube from heat, flick a few times to mix and place sample on ice. This will be your stock RNA sample. <span style="font-family: 'Times New Roman','serif';"> 25. Quantitate RNA yield using Nanodrop spectrophotometer.

<span style="font-family: 'Times New Roman','serif';">Results and conclusions: <span style="font-family: 'Times New Roman','serif';">This lab session was only preparatory and thus we do not have any results present. <span style="font-family: 'Times New Roman','serif';">Reflection: <span style="font-family: 'Times New Roman','serif';">This lab was useful to learn a few new techniques to process RNA and to understand the purpose of a primer solution. It also required high precision work with pipettes because there were minimal amounts of liquids to be handled.

Lab 1 – October 1st 2013
==== The purpose of this lab was to learn how to extract and isolate DNA and RNA from a tissue sample. In addition we learned some basic general lab safety procedures and how to use some of the lab equipment. A first important consideration is the quantity tissue sample needed for the extraction. and amount between 25-50g is needed and between 50 and 100 g for RNA. the tissue samples in our lab session came from two species of of oyster, Pacific and Olympic and were sampled from gill or mantle. ====


 * ====micropipettes (1-1000uL)====
 * ====sterile filter pipette tips (1-1000uL)====
 * ====sterile (RNase free) 1.5mL microcentrifuge tubes====
 * ====sterile disposable pestles====
 * ====vortex====
 * ====ice buckets====
 * ====gloves====
 * ====lab pens====
 * ====safety glasses====
 * ====TriReagent====

The first part of the experiment consisted in extracting the RNA and store in the freezer so that it could be used for the following lab. We learned that handling RNA requires a good amount of attention not only because it needs to stay below a certain temperature but also because it is easier to contaminate. The samples were constantly kept on ice aside from the times that they needed to be handled. We utilizing a chemical called TriReagent that allows to separate RNA from the other components of the cell. TriReagent is made of three agents that allow it to the natural rate the proteins and keep down insoluble while keeping RNA soluble. Here follows the step-by-step procedure.

in this second part of the lab session we used DNazol to precipitate DNA from cell. We precipitated DNA by the use of ethanol and subsequently make DNA soluble by using water. We also had to centrifuge sample more than once to remove the unwanted components of the cell such as RNA and insoluble tissue fragments. At the end of this process we also verified the quality of our DNA sample by using the NanoDrop. The nano drop allows to measure the DNA concentration and the ratio between A260/A280 and A260/A230. The results were as follows: A260/280 ratio=1.89, A260/230 ratio=0.43 and 151 µg/ µL concentration. Reflections the purpose of this lab was to be exposed to the basic procedures for RNA and DNA extraction and to learn how to properly handle both equipment and samples. Additionally the lab was also important to learn how to quantify the quality of a DNA/RNA extraction. The procedure followed in the lab allowed us to measure the concentration of DNA in the test tube. I found this lab extremely important not only to get acquainted with that equipment and procedures but also because being able to understand how DNA/RNA are extracted is something that every scientist should be exposed to regardless of its field of study. Also the ramifications of DNA extraction has been touching virtually every life science discipline as the information that DNA can provide is of extreme importance and relevance to many studies. I would have liked to have had a bit more insights before starting the lab on the different applications of RNA versus DNA in current research.
 * RNA ISOLATION PROTOCOL**
 * 1) Label the tube containing tissue sample with initials and date. Keep the sample stored on ice until you are ready for homogenization.
 * 2) Add 500uL of TriReagent to the 1.5mL snap cap tube containing tissue. Store on ice.
 * 3) Carefully homogenize the tissue using a disposable pestle. If the tissue is difficult to homogenize, carefully close the tube tightly and briefly vortex the sample.
 * 4) After the sample is completely homogenized, add an additional 500uL of TriReagent to the tube and close the tube tightly.
 * 5) Vortex vigorously for 15s.
 * 6) Store tissue sample to at -80ºC.
 * DNA Isolation (DNazol)**
 * Supplies and Reagents**
 * micropipettes (1-1000 µL)
 * sterile filter pipette tips (1-1000 µL)
 * 1.5 mL microfuge tubes
 * microcentrifuge tube rack
 * microcentrifuge (room temperature)
 * razor blades
 * vortexes
 * DNazol
 * 100% ethanol
 * 75% ethanol
 * 0.1% DEPC water
 * kim wipes
 * Nanodrop
 * DNazol Extraction Protocol (Adapted from MRC manual)**
 * 1) Using a sterile pestle, homogenize your tissue sample in 0.5 mL of DNazol in a 1.5 mL sterile microfuge tube. After the tissue is homogenized, add 0.5 mL more of DNazol and mix well.
 * 2) Let your sample incubate for 5 minutes at room temperature.
 * 3) Spin your sample at 10,000 x g (room temp) for 10 minutes.
 * 4) Transfer your supernatant to a new, labeled tube.
 * 5) Add 0.5 mL of 100 % ethanol to your sample.
 * 6) Mix your sample by inverting your tube 5-8 times.
 * 7) Store your sample at room temperature for 1 minute.
 * 8) Your DNA should form a cloudy precipitate. Remove the DNA and put in a new tube using your pipette.
 * 9) Let your sample sit at room temp for 1 minute and remove the rest of the lysate (liquid that is not DNA).
 * 10) Wash your DNA with 1 mL of 75% ethanol: Pipette the ethanol into your DNA tube, invert 6 times, and let sit for 1 minute. Remove the ethanol from the tube and repeat.
 * 11) If there is ethanol left at the bottom of your tube after the second wash, remove with a small pipette.
 * 12) Add 300 µL of 0.1% DEPC water to your DNA and pipette up and down multiple times to dissolve.
 * 13) Bring your DNA sample up to the Nanodrop to quantify.
 * DNA Quantification**
 * 1) Pipette 2µL of 0.1%DEPC-H20 onto the Nanodrop pedestal and lower the arm.
 * 2) Select "dsDNA" from the pulldown menu
 * 3) Click "Blank", to zero the instrument. NOTE: steps 1 and 2 only need to be done once for the whole class.
 * 4) Pipette 2µL of your DNA sample onto the Nanodrop pedestal and lower the arm
 * 5) Click "Measure". Record your DNA concentration (ng/µL), A260/280 ratio and A260/230 ratio. NOTE: The Nanodrop uses the Beer-Lambert Law to calculate DNA concentration for you.
 * 6) Raise the arm and wipe off you sample with a KimWipe
 * 7) Clearly label your stock DNA sample with the word "DNA", source organism/tissue, your initials, today's date and the concentration in ug/uL.
 * 8) Store sample at -20ºC.