Friedman Lab Protocols



Hazardous Waste Protocol

  1. Place labeled waste container in ziplock bag
  2. Mark hazardous waste label (found on the Wiki forms page) with the following:
    1. Chemical composition percentage (Note: Must add up to 100%!)
    2. Use check boxes to note any associated hazards. If other please explain.
    3. Labeled by
    4. Department
    5. Phone
    6. Building
    7. Room
Hazardous_Waste_label_example.jpg


  1. Attach label to outside of ziplock bag that the hazardous waste is in with tape. (Note do not use staples as this could puncture the bag!)
  2. Notify Sammi via email that there is waste that needs to be picked up and she will contact EH&S. Please make sure to include the volume of waste, number of containers, type of container, and location of waste in the email. (Note: if waste needs to remain in hood please keep in the hood. EH&S will remove from there)

EH&S will remove waste 2-4 weeks after chemical waste collection request is submitted.

Thermalcycler program: 16S-GRAD

Thermocycler used is MJ Research 98-well ThermalCycler.
RME August 2001
This program may be used to determine the optimal annealing temperature for the amplification of 16S rDNA.
Thermocycler Program “16S-GRAD”
1) 94°C 00:03:00
2) 94°C 00:01:00
3) 40-52°C gradient across 12 columns of wells *
4) 72°C 00:02:00 +5 sec/cycle
5) to step 2, 28 times
6) 94°C 00:01:00
7) 50°C 00:01:30
8) 72°C 00:07:00
9) 4°C 00:00:00
10) END
* Annealing Temperature Gradient
Column (L to R)
Temperature °C
1
40.0
2
40.3
3
40.9
4
42.0
5
43.4
6
45.1
7
47.1
8
48.8
9
50.2
10
51.1
11
51.8
12
52.0

Abalone Fungal Lesion Collection Kit

This kit contains a histo-pen, cryovials, ziplock bags, fungal media plates, ethanol burner, filtered seawater with antibiotic (penn-strep), forceps, scalpel, parafilm, 1% iodine, transfer pipets, and a beaker.
For all abalone shell lesion collection:
  1. Rinse lesion with 70% ethanol.
  2. Rinse with 1% betadine.
  3. Rinse with filtered seawater.
Collect 5 lesions for storage in ethanol.
  1. Place lesion in cryovial tube.
  2. Fill tube almost to the top with 70% ethanol.
  3. Seal tube with parafilm.
  4. Label tube with contents. (“White abalone shell, fungal lesion, 70% ethanol.”)
Collect 5-15 lesions on agar fungal media plates.
  1. Stab a piece of lesion halfway int to agar.
  2. Place up to three lesions on the plates, spread apart.
  3. Label the collection date on the plate.
  4. Seal the plate with parafilm.
Store tubes and plates in a refrigerator or cooler until it gets back to the UW. Keep them double bagged.

Big Gel(R. Estes, 11/03)
1. Add approximately 0.8 g agarose (molecular grade) to 100 ml of 1x TBE buffer in a 200 ml flask. [Always use the ethidium bromide contaminated flask in the gel area.]
2. Cover with Saran wrap and poke holes in the top. Heat the flask in the microwave and swirl periodically, watching that it does not boil over. Heat just long enough for all the agarose to dissolve, about 3 minutes. You can tell when the agarose is dissolved when there are no translucent “lenses” or “disks” floating around.
3. Allow to cool until just cool enough to touch without instantly burning your hands. Add 10 ml of ethidium bromide stock solution (1 ml/10 ml) to the molten agarose. Swirl.
4. Pour cooled yet molten gel into gel box. Add the appropriate combs to form wells.
5. After the gel is solid, orient the gel so that the wells are toward the – (black) electrode. The DNA should run towards the + (red) electrode. [The gel is solid after it turns opaque. This takes about 30 minutes.]
6. Pour TBE buffer over the gel so that it is covered by 1 cm of 1x TBE buffer solution. Gently remove the combs.
7. Load ladder and samples on gel by pipetting the following mixtures into the wells. Try not to touch the side of the wells with the pipet tips as this can warp the DNA bands. [See notes on gel loading.]
Ladder: 1 ml loading dye Samples: 5 ml PCR product
1 ml DNA ladder 1 ml loading dye
4 ml npH2O
8. Hook up the gel to the power box and run the gel at 130 volts until the dye front is almost to the edge of the gel. Running at lower voltage gives more precise bands but takes longer.
9. After running the gel. Rinse out flask for gel preparation in the stainless steel sink with lots and lots of water. Dry on rack in gel area.
10. Reuse electrophoresis buffer until it gets blue from the loading dye. If samples start to float away when loading, add 2 ml of loading dye to sample. This should help the sample sink. This is a sign that the electrophoresis buffer needs to be changed.
11. Store used electrophoresis buffer in the gel rig with the lid on. This will help it keep from evaporating.

Extraction of DNA from Celite Beads for PCR Amplification

(Jennifer L. McLarnan 11/99, revised from “Extraction of DNA from Sediments”, James P. Gray 5/96)
Important: This protocol assumes the celite sample contains NO sediment. If sediment is included see the protocol “Extraction of DNA from Sediments for PCR Amplification.”

Extraction

1) Make lysis buffer and heat at 55C for 60 minutes. Lysis buffer must be made just prior to use and heated for 60 minutes before using. Use buffer warm.
Buffer: -500mM NaCl (100ul of 5M per ml)
-50mM EDTA (100ul of 0.5M per ml)
-50mM Tris pH8.0 (50ul of 1M per ml)
-4% SDS (160ul of 25% per ml)
-npH20 (590ul per ml)
2) Into a 2.0ml beadbeater tube weigh out the following (in order):
2.0g 0.10mm ceramic beads
200.0mg celite bead sample
900.0ul lysis buffer (55C)
3) Homogenize samples in beadbeater for 1.0min on the “homogenize” setting.
4) Incubate tubes in 70C waterbath for 60 min.
5) Centrifuge samples at 13,000rpm for 2 minutes using the Eppendorf 5415C centrifuge.
6) Transfer supernatant to a new 1.7ml tube. Incubate samples 15.0min on ice.
7) Centrifuge samples in the 5415C centrifuge that has been pre-chilled to 4C. Centrifuge at 14,000rpm for 5.0min at 4C.
8) Transfer supernatant to a new 1.7ml tube and add 2 volumes of –20C 100% ethanol.
9) Invert tubes 20 times.
10) Incubate tubes at –20C for 15minutes.
11) Centrifuge samples at 14,000rpm (4C) for 5.0 minutes.
12) Discard supernatant and wash pellets once with 1.0ml –20C 70% ethanol.
13) Air dry pellets.
14) Resuspend in 50ul TE (pH 7.5).
15) Proceed immediately to Purification.

Purification

1) Quantitate the extracted DNA using the GeneQuant II.
2) Purify samples via Bio 101 GeneClean III Kit with Spin. Follow directions for purification from solution. For each column, add only 10ug of DNA and bring the DNA volume up to 100ul with npH2O. Bind for 20 minutes at room temperature with vortexer set at constant shake, speed 2. After wash steps, centrifuge empty columns 15 minutes at 14,000rpm to dry pellets. Elute samples once using GeneClean Elution Solution, incubate at room temperature for 10 minutes (with occasional flicking) prior to centrifuging.
3) Run the samples on a 1% Agarose gel (1x modified TAE) at 5 V/cm.
4) Excise the large MW band using a clean razor blade for each sample.
5) Isolate the DNA from the agarose using Bio 101 GeneClean III Kit with Spin. Follow directions for purification from agarose.
6) Quantitate the DNA using the GeneQuant II.
7) Proceed immediately to PCR Amplification.

PCR Amplification

1) Set up PCR reactions per separate protocol.
2) Run thermocycler program “16S-OLD” on the MJ Research Minicycler.
3) After cycle is complete verify product by running 5ul on a 0.5% Agarose gel (1xTBE).

Chemical Spill Basics

Hazardous material spills that do not endanger workers in the immediate area may be cleaned up by area personnel who have been trained and are properly equipped to clean up the spilled material safely. **Spill kits** are available from University Stores during business hours.
Hazardous material spills that cannot be safely adsorbed, neutralized, or otherwise controlled at the time of release by employees in the immediate release area are considered to be emergencies requiring outside assistance by the Seattle Fire Department (SFD), Environmental Health & Safety (EH&S), and possibly a spill cleanup contractor.
When in doubt about whether you need help or not, it is best to call for help. EH&S staff cannot clean up spills but can offer advice on how to handle spills. Call 206.543.0467.
When you need emergency help, do the following:
  • Alert others and evacuate all affected areas.
  • Pull the fire alarm if needed or report the incident to UW police:
·
    • UW Campus or UWMC dial 9-911
    • Harborview dial 3000
The UW Police will notify the Seattle Fire Department (SFD) who will respond, stabilize, and contain the spill. Environmental Health & Safety (206.543.0467) will advise SFD as needed. The incident may require use of a spill clean up contractor at the department's expense. All waste must be **contained and labeled** as instructed by EH&S.

16S rDNA Sequencing Outline

· Isolation of bacterial DNA using Instagene
· PCR amplification of 16S rDNA
· Cloning and Transformation using E.coli cells
· Isolation of plasmid DNA using an alkaline lysis mini-prep
· Confirmation of 16S rDNA insert by PCR and restriction digest
· 16S rDNA Sequencing

Isolation of Bacterial DNA using Instagene

(Ben Paulson- 4/2/03)

Required equipment: 56 °C water bath, 100 °C heat block
  1. Plate desired isolate under appropriate growth conditions.
  2. Select one, or a few, colonies and transfer them using a sterile loop to 1 mL dH2O in a 1.7 mL microfuge tube.
  3. Centrifuge for 1 minute at 12,000 rpm and remove supernatant.
  4. Briefly mix Instagene matrix on a stir plate.
  5. Resuspend pellet with 200 μL Instagene matrix (use a P1000 tip to transfer matrix solution).
  6. Incubate at 56°C for 15-30 minutes.
  7. Vortex at high speed for 10 seconds.
  8. Place tube in 100 °C heat block for 8 minutes.
  9. Vortex at high speed for 10 seconds.
  10. Centrifuge at 12,000 rpm for 3 minutes.
  11. Use immediately or store at –20 °C.
  12. When reusing, repeat steps 9 and 10.

PCR Amplification of 16S rDNA

(Jennifer McLarnan, rev. by Ben Paulson 4/3/03)

  1. Thaw all components of cocktail (see below), except for Taq polymerase at room temperature. Leave Taq polymerase in freezer.
  2. Once thawed, briefly centrifuge and place components on ice.
  3. Label PCR tubes and place on ice.
  4. Prepare a PCR reaction cocktail on ice as described below. This outline is designed for 20 μL reactions. Each of the following values is per reaction.
Component
Volume (μL)
10x Mg2+ free buffer
2.0
25 mM MgCl2
1.6
50% Acetamide
2.0
PCR H2O
8.6
25 mM dNTP mix
0.16
50 μM Forward Primer (8fm)
0.2
50 μM Reverse Primer (1492RU-rph)
0.2
Taq Polymerase
0.24
Total
15
  1. Into each of the PCR tubes place 2 μL PCR H2O and 3 μL sample. For a negative control use 5 μL PCR H2O and no sample.
  2. Aliquot 15 μL of the cocktail into each of the sample tubes. Mix by pipetting up and down and change tips between each sample tube.
  3. Add 5 μL Pink Wax to each tube.
  4. Place PCR tubes in thermocycler and run program “16S-JLM.”

Cloning and Transformation of 16S rDNA
(R. Estes, 12/03)
Required equipment: 42 °C water bath, LB plates with 50-100 μg/mL ampicillin (two per transformation), 37 °C incubator with shaker.
  1. Equilibrate water bath to 42 °C.
  2. Warm SOC medium (found in –80 °C freezer) to room temperature.
  3. Thaw on ice 1 vial of One Shot Chemically Competent E. coli cells for each transformation.
  4. Using PCR tubes set up the following cloning reaction:
Reagent
Volume (μL)
PCR Product
3
Salt solution
1
Sterile H2O
--
TOPO Vector
1
Total Volume
5
  1. Mix each reaction gently and incubate for 30 minutes at room temperature.
  2. This reaction mixture may be stored at –20 °C overnight or placed on ice to proceed to transformation.
  3. Add 2 μL of the TOPO cloning reaction to a vial of One Shot Chemically Competent E. coli cells and mix gently (no pipetting).
  4. Incubate for 5 minutes on ice.
  5. Heat shock cells for 30 seconds in 42 °C water bath.
  6. Transfer tubes to ice.
  7. Add 250 μL room temperature SOC medium to each tube.
  8. Shake for one hour at 37 °C (200 rpm).
  9. Place LB-Ampicillin plates in 37 °C incubator for approximately 30 minutes.
  10. Plate 75 μL on LB-Ampicillin plates.
  11. Grow 24-48 hours at 37 °C.

Isolation of Plasmid DNA using an Alkaline Lysis Mini-Prep

(Ben Paulson- 4/3/03)

  1. Inoculate 1-3 mL LB-Ampicillin broth with a single transformed colony.
  2. Incubate overnight at 37 °C with shaking.
  3. If needed, transfer broth culture to 1.7 mL microfuge tube.
  4. Centrifuge at 8000 rpm on microcentrifuge for 10 minutes. Discard supernatant.
  5. Resuspend each pellet with 200 μL GTE solution. Incubate for 5 minutes at room temperature.
  6. Add 400 μL of freshly prepared lysis solution. Mix gently and incubate on ice for 5 minutes.
  7. Add 300 μL of ice-cold 3M potassium acetate. Mix gently and incubate on ice from 5 minutes.
  8. Centrifuge for 1 minute at 8000 rpm at 4 °C.
  9. Transfer supernatant to a clean microfuge tube being careful not to transfer any pellet material. Multiple centrifugations may be required to eliminate pellet transfer.
  10. Add 540 μL isopropanol to supernatant and incubate at room temperature for 2 minutes.
  11. Centrifuge for 1 minute at 14,000 rpm to pellet the plasmid DNA. Discard supernatant.
  12. Wash the pellet twice with 1 mL 70% ethanol. Centrifuge for 5 minutes at 8000 rpm and discard supernatant.
  13. Let pellet air dry (overnight).
  14. Resuspend with 50 μL PCR H2O or TE pH 7.5. Add RNaseA if necessary and incubate at room temperature for 30 minutes.
  15. Store at –20 °C until use.
  16. Presence of 16S rDNA insert should be confirmed by PCR/restriction digest prior to sequencing.

16S rDNA Sequencing (From Plasmid Preps)
(R. Estes, 12/03)

We complete our sequencing through the Marine Molecular Biotechnology Lab (MMBL) located in the Marine Studies Building. The current sequencing technician at MMBL is Ann Riddle (ariddle@u.washington.edu). Any questions should be directed through her. Sequencing is done using the DYEnamic ET Dye Terminator Cycle Sequencing Kit and samples are analyzed with a MegaBACE 1000 system. When you are ready to begin sequencing, contact Ann and she will provide you with Sequencing Premix, 7.5 M ammonium acetate, and sequencing instructions. A copy of these sequencing guidelines is provided along with this SOP.

Analysis of Plamid Concentration using GeneQuant

1. Turn on GeneQuant and printer.
2. Obtain quartz cuvette from drawer labeled “GeneQuant Supplies and Manuals”
3. Place 70 μL dH2O into the cuvette and position it above the insertion point of the machine.
4. Press “Set Ref” on the machine and wait for machine to instruct you to insert cuvette. Remove cuvette when prompted.
5. Remove dH2O from cuvette and replace with 68 μL dH2O and 2 μL plasmid prep solution.
6. Position cuvette above insertion point and press “Sample.” Again, insert and remove sample when prompted.
7. Following sample reading, press “Abs,” “Ratio,” “Conc,” and “Protein.” These will print out the values for absorbance at 280 nm, ratio of A260/A280, dsDNA concentration, and protein concentration respectively.
8. Using the value for dsDNA concentration determine the volume necessary to add ~200 ng plasmid to each sequencing reaction.
*Note- the amount of plasmid added seems to be variable. One can try different values to obtain best results.*

Sequencing Reactions

*Note- The sequencing pre-mix contains dye molecules that are light sensitive. Care should be taken to avoid excessive exposure to light throughout the sequencing process.*
  1. Determine the primers to be used for sequencing and calculate the volume to add to each reaction so that 5-10 pmol primer are added.
  2. Each sequencing reaction should contain the following:
· ~200 ng plasmid
· 5-10 pmol primer
· 4 μL sequencing pre-mix
· dH2O to a final volume of 10 μL
  1. Sequencing reactions are run on the thermocycler using the “Seq-30” program:
40 cycles of : 95 °C for 20 s
50 °C for 15 s
60 °C for 60 s
  1. Sequencing reactions may be stored at 4 °C for a short period of time before clean up, but should be kept out of the light.

Ethanol Precipitation for DNA Sequencing (Clean-up reaction)

*Keep samples in the dark as much as possible*
The Friedman lab has generously allowed us to use their large centrifuge for this procedure.
1. Set centrifuge to 4 °C.
2. Mix 7.5 M ammonium acetate with 100% ethanol according to the following:
1 μL ammonium acetate/reaction
27.5 μL 100% ethanol/reaction
3. Add 28.5 μL of the mix from step 2 to each sequencing reaction and vortex.
4. Centrifuge samples at 3250 rpm for 30 minutes at 4 °C.
5. With 10 minutes left in spin turn centrifuge temperature back to 25 °C.
6. Do not let samples sit for more the 5 minutes following centrifugation or DNA may resuspend.
7. Remove lids from samples and cover openings with several paper towels. Invert samples and gently shake out liquid. *Do not bang on counter*
8. Cover openings with new dry paper towels and load samples into centrifuge upside down (for inverted spin).
9. Centrifuge for 3 minutes at 600 rpm (keep rpm’s near this value).
10. After inverted spin check that samples are dry and repeat spin if necessary.
11. When samples are dry add 200 μL 70% ethanol to each sample. Place lids back on.
12. Centrifuge samples at 3250 rpm for 3 minutes (room temperature).
13. Repeat step 7
14. Do an inverted spin with dry paper towels for 5 minutes at 600 rpm.
15. Repeat step 10.
16. Place samples under hood for 20-25 minutes.
17. Place lids on samples and store at –20 °C until sequencing is to be completed.
When samples are ready to be sequenced, contact Ann and let her know the number of samples you have. She will let you know when to bring them by the MMBL lab (have her tell you where to put them). Along with the samples you should leave a PC-formatted zip disk and a list of the sequencing reactions (i.e. 1-16, or BP1-BP16). Make sure that PCR tubes are labeled accordingly.

Deparaffinization

(R. Estes Strenge, 5/07)
Use coplin staining jars under the hood for this process.
1. Wash slides in HistoClear 3 x 3 min.
2. Wash in 100% EtOH 2 x 3 min.
3. Wash in 95% EtOH for 3 min.
4. Wash in 80% EtOH for 3 min.
5. Wash in 70% EtOH for 3 min.
6. Wash in 50% EtOH for 3 min.
7. Wash in 30% EtOH for 3 min.
8. Wash in PBS (pH 8.0) for 3 min.

Recipe for Ethanol
Ethanol (ml)
dH2O (ml)
TV (ml)
% EtOH
60
0
60
100%
57
3
60
95%
48
12
60
80%
42
18
60
70%
30
30
60
50%
18
42
60
30%

DNA Quanting via Standards in Layout

Updated: 5/22/2013 SJW

  1. Turn on plate reader. Allow at least 20 minutes for lamp to warm up before use.
  2. UV a clean plate for 3-5 minutes.
  3. Thaw PICO green and prepare enough of a 1:200 solution of the PICO green for the number of wells you will be loading. You will need 50ul per well. Make sure the PICO is fully thawed and mixed. e.g.- If you are going to have 20 wells then, 20 X 50=1000. 1000/200=5 (therefore you would add 5ul of PICO stock to 995 1x TE buffer). Make at least 10% more than you will need so that you don’t run out. Remember to account for standards and blanks. All samples, standards and blanks should be run in duplicate.
  4. First add 49ul of TE to each well you will use, including the standards and BLANKS.
  5. Get the DNA Quanting standards out of the fridge in the box labeled (“DNA Quanting buffers). There are a 100, 80, 60, 40 , 20, 10, and 5 ng/ul standards.
  6. Pipette 1ul of your DNA template to each of your unknown wells. Then, pipette 1ul of the appropriate standards to the appropriate wells. Lastly, pipette 1ul of TE into each BLANK well.
  7. Pipette 50ul of your 1:200 PICO solution on top of each of the wells; pipette up and down to mix.
  8. On the computer open Magellan.
  9. Put your plate in and run the method called “DNA Quanting via standards in layout”.
  10. After the plate has run/Analysis… Click “View data” button. The first thing you want to look at is the standard curve graph under the “Concentration” tab. You should have an R-squared value of greater than 95%. If your R squared is less than 95% or any of your points have a large standard deviation then look into the troubleshooting at the end of this protocol.
  11. If the standard curve looks good then your unknowns should have quanted well. Under the “Concentration” tab click on the “Single Concentration tab” to check the quant of your unknowns and especially check your Quant-it standard(Q). Your Quant-it® standard should have quanted almost right on if your standard curve is correct.

Notes

There are 50ul aliquots of PICO green in the 236 (-20° C).
If your samples quant over 100ng/ul they will appear with a # sign in front of them (e.g. # 102.06). This simply means that your data is based on an extrapolation factor. It does not mean that it is incorrect data or anything.
Notice that the BLANKS are in the last vertical row not the last horizontal row as before!

Troubleshooting

If your standard curve is off……. You can either run another plate… or click on the “Transformation tab” under the view data screen and then click on the “blank reduction” tab. Look at the fluorescence values it gives for your standards. It may be that just one of your rows of standards is off. You can check this by copying these blank reduction values into Excel and plotting the rows individually into graphs and checking the R squared value of each row. You may find that one of your rows is bad (low Rsquared) and that one is good (high R squared). If this is the case you want to go into the “edit currently run method” mode and change the bad row into just samples and therefore just use the good row as your standard curve.

Agarose Gel Electrophoresis(R. Estes, 11/03)
Agarose gel electrophoresis is a method used to separate DNA fragments with an electric current. The basic idea is that a solid support (the gel) is cast on a single plate (agarose). The slab is placed between electrode compartments with the polarity of electrodes depending on the material being separated. A small sample of the material being separated is placed in each preformed well.
High-density material is added to the sample with a tracking dye to ensure that the material stays in the well until drawn into the gel itself. Loading dye is required because agarose electrophoresis is a “submarine” techniques-->
Gel loading dye serves three purposes in DNA electrophoresis: First, it increases the density of the sample, ensuring that the DNA will drop evenly into the well. It adds color to the sample to simplify loading. Finally, the dyes migrate in an electric field towards the anode at predictable rates which enables one to monitor the electrophoretic process. To increase the sharpness of DNA bands, we use Ficoll (type 400) polymer as a sinking agent instead of glycerol. The use of the lower molecular weight glycerol in the loading buffer allows DNA to stream up the sides of the well before electrophoresis has begun and can result in a U-shaped band. In TBE gels, glycerol also interacts with borate which can alter the local pH.

We use 0.8-1.0% agarose gel for most applications. The greater the percentage of the gel, the less mobility will be shown by the molecules. The gel should be about 0.5 cm thick and covered with approximately 1 cm of TBE buffer. When separating DNA in agarose, the + (red) electrode should be at the bottom and the – (black) at the top, since this will then cause the DNA to migrate towards the + (red) pole and therefore through the gel. The fragments are then visualized with ethidium bromide as the staining agent, which fluoresces brightly when stimulated with short-wave UV, however one can also use other fluorescent dyes.

Safety IssuesEthidium bromide is a mutagen that binds DNA. It is best to assume that everything having to do with running gels is contaminated with ethidium bromide. This includes gel boxes, anything in gel room including sink, cabinets, camera system. It also includes the gel pouring area and microwave. Always wear gloves and lab coat while working with potentially contaminated equipment. Do not touch the rest of the lab with potentially contaminated gloves. Remove gloves immediately when done and wash hands.

Gel Loading

(R. Estes 11/03)
It is important to load your gel in a way that is easy for you and others to understand. While you might know what you are looking at right after you ran it, trying to figure out poorly organized gels when going over your lab notebook months later can a chore! Keep in mind that the PI (primary investigator, ie. Carolyn) and other lab personnel need to be able to read your lab notebooks, even when you are not here.
Have your gel labeled as soon as possible in your lab notebook. It is preferable that you have this ready BEFORE showing your gel to the PI. Unless you have a very simple gel and you know what is in each lane, do not take it to a supervisor to look at. If you do not know what is on your gel, nobody else will either.
Keep these points in mind when making and labeling gels:
  • ALWAYS run a negative PCR control. [PCR cocktail and npH2O ran through thermal cycler with samples.]
  • ALWAYS run a positive control. [If you know that some amples are positive, this will do.]
  • Run a DNA ladder in at least the far left well of each row. You may want to run a ladder lane on each side of the row. DNA ladder is essential to knowing what our band sizes are. Run gels long enough to see separation in ladder bands.
  • When taking a picture, make it as nice and easy to read. Zoom in on the gel to fit as much of it in the photograph as possible.
  • Label each lane used on a gel.
  • Run samples in a logical order. [Chronological by sample number, by date, concentration, etc.
  • In your lab notebook, have a table that explains what is in each lane on your gel.
  • When en electronic gel is requested, that means that the PI wants to send it to someone or place it in a manuscript
Example. Jane has seven OHV samples, ran in duplicate, and in three different concentrations (400, 600, 800 ng). She makes sure to have all samples being tested at each DNA concentration on the same row, the samples lined up horizontally according to chronological sample number, and the samples that are the same lined up vertically. This makes it easier for everyone to look at her gels. Way to go Jane! J


Jane’s Gel
gel_ex.png

Jane’s Gel










ng DNA
Lane
1
2-3
4-5
6-7
8-9
10-11
12-13
14-15
16

Sample
1Kb Ladder
9-13
10-3
10-5
11-14
13-6
13-9
13-13
Neg Control
400
Lane
17
18-19
20-21
22-23
24-25
26-27
28-29
30-31


Sample
1Kb Ladder
9-13
10-3
10-5
11-14
13-6
13-9
13-13

600
Lane
32
33-34
35-36
37-38
39-40
41-42
43-44
45-46


Sample
1Kb Ladder
9-13
10-3
10-5
11-14
13-6
13-9
13-13

800
Jane lists her samples in her notebook like this.
Lane
Sample
ng DNA
1
1Kb Ladder

2-3
9-13
400
4-5
10-3
400
6-7
10-5
400
8-9
11-14
400
10-11
13-6
400
12-13
13-9
400
14-15
13-13
400
16
Neg Control

17
1Kb Ladder

18-19
9-13
500
20-21
10-3
500
22-23
10-5
500
24-25
11-14
500
26-27
13-6
500
28-29
13-9
500
30-31
13-13
500
32
1Kb Ladder

33-34
9-13
600
35-36
10-3
600
37-38
10-5
600
39-40
11-14
600
41-42
13-6
600
43-44
13-9
600
45-46
13-13
600

General Cloning Method (16s Genes)

(Robyn M. Estes 7/28/00)

PCR

1) Use universal primers with cloning restriction enzyme sites (8fb-JP and 1492RU-rph) for 16s rRNA amplification w/ links for directional cloning.
2) Run on a 1% Seakem agarose gel with 1X TBE and 1ml/ml EthBr to make sure it worked.
3) Purify the PCR product using a Bio Gel P30 column.
4) Quantitate the PCR product and the pNEB from the freezer using the GeneQuant II.

DNA Digest

5) Calculate 10mg DNA (PCR product or pNEB) per reaction.
Cocktail: 10X NEB Buffer #4 5
100X NEB BSA 0.5
Enzyme: Pac I 2
Asc I 2
DNA template 10mg
npH2O ?
Total volume 50ml
6) Incubate in 37C water bath overnight.
Gel purification: 1X modified TAE, 1% SeaPlaque, 5V/cm.
Add guanosine (same way as EthBr) to the gel and buffer.
Post-stain the gel with ethidium bromide.

Ligation

7) Excise the bands using a clean razor blade for each sample.
Go fast and set the UV @ 70%.
8) Melt the gel slices in the 70C water bath for about 15 minutes.
9) Combine the gel slices (pNEB + insert + H2O) for a TV of 9ml.
1:3 molar ratio = 100ng pNEB : 165ng insert
1:1 molar ratio = 100ng pNEB : 55ng insert
10) Place in 37C water bath for 5 minutes.
11) Add 11ml of ice cold cocktail for a TV of 20ml.
Cocktail: 10X ligase buffer 2
T4 DNA ligase 2
npH2O 7
12) Mix and place on ice.
13) Incubate @ 4C overnight.
14) Freeze for 30 minutes then centrifuge on high for 15 minutes @ 4C.

Transformation

15) Slowly defrost cells (Stratagene XLI Blue MRF¢ - Epicurian coli) on ice from –80C.
16) Mix 2ml of ligation product with 40ml cells in an eppendorph tube and place on ice.
17) Place the cuvettes on ice.
18) Put the cells in the cuvette and electroporate w/ 1 pulse @ 1.7kV.
19) Immediately resuspend the cells with 1.0 ml SOC broth.
20) Transfer the cells to sterile tubes and incubate @ 37C for 60 minutes and set the shaker @ 250rpm.
21) Overlay the LB-Amp plates w/ 100ml each of X-gal and IPTG 30 minutes b/f plating.
22) Spread 100ml of cells onto plates and incubate @ 37C overnight.
23) Controls: 2ml pNEB electroporated into cells (should see TNTC blue colonies)
Spread plain cells on plate w/o electroporation (should see NG)
2ml pUC18 electroporated into cells (should see approx. 500 blue colonies)
(5ml pUC18 + 100ml SOC on plate)

Genotyping on MegaBACE

Sample preparation

PCR on samples with microsatellite primers.
Add PCR products to fresh PCR plate, add npH2O to get 10 ml of total volume.
(May need optimizing with different amounts of each PCR product.)
Follow MMBL protocol for “Ethanol Precipitation for DNA Sequencing”.
Store samples at -20°C until ready to resuspend.
Resuspend PCR products with 4 npH2O (up to 24 hours prior to running samples).
Add 2 ml of resuspended PCR products to MegaBACE plate.
Dilute 0.2 ml of MegaBACE size standard in 2.8 ml loading solution for each sample.
(96-well plate: add 21 ml of size standard and 294 ml loading solution.)
Add 3 ml of size standard/loading solution mixture to each well.
Total volume for each well should be 5 ml. For unused wells add 5 ml loading solution.

Running samples

Turn on hotplate for denaturization.
Wake up the MegaBACE 2-3 hours b/f your run.
MegaBACE is awake at 44°C for up to 4 hours. (Longer will damage capillaries.)
MegaBACE is asleep at 25°C for up to 72 hours.
Leave the control manager open on the computer.
Chose one of two modes on the control manager: “Sequencing” and “Genotyping”.
Configure: Select Application: Genotyping.
Ignore windows pop-up.
“Rinse Tips” at start of day only.
Have two buffer plates (filled with 200 ml 1X LPA buffer) ready to go in green plates.
Use one old and one new, OK to use two new.
Have sample ready (5 ml of DNA/Ladder/Loading Solution mixture) in white plates.
Spin down 6 matrix tubes (stored in fridge) for 2 min at 3000 RPM.
Placed use tubes in rack beside MegaBACE to desiccate.
Cover and spin new buffer plate and sample plateup to 3000 RPM then stop.

Immunofluorescent Antibody Test (IFAT) Protocol for Vibrio parahaemolyticus

(R. Estes Strenge, 5/07)
1. Using a Pap pen, create a well around the site on the slide to be analyzed.
2. Wash slides in PBS 3 x 10 min.
3. Block slides in 100 ml PBST + 2% BSA for 30 min.
4. Rinse off with 200 ml PBST + 1% BSA.
5. Incubate slides with 1° antibody for 1 hour in humidity chamber using PBST + 1% BSA as the diluent. Dilute at 1:25 ratio (each slide = 2 ml 1° / 50 ml diluent).
6. Wash in large volume (700 ml per 5 slides) of PBST 3 x 10 min.
7. In dim light: Incubate in FITC conjugated 2° antibody* for 30 min with PBST + 1% BSA as the diluent. Dilute at 1:80 ratio (each slide = 1.25 ml 2° / 100 ml diluent).
8. Wash in large volume (700 ml per 5 slides) of PBST 3 x 10 min.
9. Flood with Evan’s blue (0.01% in PBS) for 2 min.
10. Wash slides in PBS 3 x 10 min.
11. Add a drop of Dabco + glycerol and coverslip.
12. Store slides in humid chamber at -20 °C and cover with foil.

Fixing Bacteria Slides

1. Grow up Vp overnight at room Temperature in 1-2 ml of T1N2 media.
2. Next day, spin down the cells and removed the supernatant.
3. Add 1 ml dH2O to resuspend the cells.
4. Drop 50-100 ml per slide.
5. Allow slides to air dry.
6. Pass slides quickly through a flame to help cells adhere to the slide.
7. Fix slides in MeOH for 5 min.
8. Stand up slides and allow to air dry.
9. Store slides in refrigerator in a slide box. Place the box in a ziplock bag with Dririte.


Products


1° Antibody
Vibrio parahaemolyticus, serotype 04, Rabbit antibody. Accurate Chemicals YCC310-471. (Takes 2-3 months to produce)

2° Antibody
Monoclonal anti-rabbit IgG, FITC Conjugate, Sigma F 4151 * or Alexa Fluor 488, Invitrogen A-11008

Isolation of Bacterial DNA using Instagene
(R. Estes, 12/03)

Required equipment: 56 °C water bath, 100 °C heat block
  1. Plate desired isolate under appropriate growth conditions.
  2. Select one, or a few, colonies and transfer them using a sterile loop to 1 mL dH2O in a 1.7 mL microfuge tube.
  3. Centrifuge for 1 minute at 12,000 rpm and remove supernatant.
  4. Briefly mix Instagene matrix on a stir plate.
  5. Resuspend pellet with 200 μL Instagene matrix (use a P1000 tip to transfer matrix solution).
  6. Incubate at 56°C for 15-30 minutes.
  7. Vortex at high speed for 10 seconds.
  8. Place tube in 100 °C heat block for 8 minutes.
  9. Vortex at high speed for 10 seconds.
  10. Centrifuge at 12,000 rpm for 3 minutes.
  11. Use immediately or store at –20 °C.
  12. When reusing, repeat steps 9 and 10.

Abalone Care


Routine Checks

1. Check UV, pumps, chillers are ON. If something is not working, check fuse. May need to be reset
2. Check and record water and air temp
3. Check pH – Don’t stick tubes in water! Use transfer pipette and fill to 40mL line. Rinse pH probe with DI between samples
· If pH <8.0, ADD Sodium Bicarbonate solution (stir solution and add ~1/3 scoop per sump and ½ scoop for pet & red)
· If pH >8.5, do water change
4. Check Ammonia (every other day)
· If NH3 >0.1, do water change
5. Check for MORTS – wash hands in iodine and rinse well in tap, then DI between EACH BUCKET. If you have a mort (ie: not attaching, shrunken, or dead) call Lisa (206-708-9241), Sammi (425-286-5883), or Carolyn (707-328-8388). If you call Carolyn and get voicemail do not leave a message, instead try texting her
6. Empty floor drains, check bleach tablets, and fill with iodine when full
7. Check that floor mat near door is full with dilute iodine.

Water Changes

1. Pull out the hose attached to the pump in the tank and empty ~1/2 of tank water
2. Refill tanks to appropriate level with fresh seawater located in the 2 large holding tanks near concrete beam.
5. Once drums are filled with waste water add 1 whole gallon of bleach per drum and label with date treated.
6. If you need to empty waste water drums, use pump coiled on in bucket under back sink and drain into floor
*If you have issues with filters or pumps ask and/or call Lisa or Sammi. THANKS FOR YOUR HELP!!

In-situ protocol: Antonio, 2001

(updated 7/29/09)

Day 0

Tris buffer, 1x SSC, 2x SSC, 20x SSC, Buffer 1, and Buffer 2 can all be made a day in advance.
Day 1

PCR for RLP from Abalone


Reagents
50 ml reaction (recommend 25 ul Rxn)
sdw 24.6 ml MgCl 1.5 Buffer 5 TMAC 5 DNTPs 4 BSA 10 mg/ml 0.5 Primers RA5-1 2 RA3-6 2 Taq 0.2 DNA template 3

Cycling conditions
1 = 95°C for 5:00 2 = 94°C for 1:00 3 = 50°C for 0:30 (62°C) 4 = 72°C for 0:30 5 = Go to 2, 39 times 6 = 72°C for 7:00 7 = 16°C forever

Labeling by PCR using PCR DIG Probe Synthesis Kit
Roche Applied Science: Cat No. 11 636 090 910 Add the following components to a sterile microfuge tube on ice:
Reagents
Volumes for 50ul
Volumes for 25 ul
SDW
29.25
14.625
PCR buffer (vial 3)
5
2.5
PCR DIG mix (vial 2)
5
2.5
Primers RA 5-1 (20pml/ul)
2.5
1.25
Primers RA 3-6 (20pml/ul)
2.5
1.25
Enzyme mix (vial 1)
0.75
0.375
Template DNA (1:100 dilution of RLP plasmid)
5 of e5
2.5 of e5
Note: Conditions for labeling are the same as conventional PCR outline above. Always include one reaction without DIG labeling (i.e. regular PCR reaction containing RLP DNA template). Important: Before using the DIG labeled probe, denature the probe at 95C for 3 minutes and immediately place on ice for about 30 min to separate the double stranded DNA. Store at –20 or –70C until use. 1. Set oven to 37 *C and preheat the Proteinase K in Tris-Buffer in a slide case. (For 30 mL of solution, place 0.075 mL of Pro-K in solution) 2. Prepare Prehybridization buffer and store in 4 *C fridge


Tissue Deparaffinization

~ 1 hr
1. Deparaffinize with 3 changes of SafeClear for 10 min each. (30 min)
2. Hydrate with graded ethanol series of (100,100, 95, 80, 70, and 50%), for 3 min each. (18 min.)
3. Then rinse slides in sterile dH20.
4. Equilibrate tissue sections in Tris buffer pH 7.2 for 5 min.

Permeabilization of tissues

~ 1.20 hr
1. Add tissues with 50 ug/ml Proteinase K in Tris-buffer for 45 min at 37 oC.
2. Rinse tissues with PBS: 3x10 min. (30 min).

Prehybridization

~ 1-2 hr
1. Prepare prehybridization buffer (Total volume = 1 mL): 0.51 mL deionized formamide 0.20 mL 20x SSC 0.05 mL heat-denatured sperm DNA (10 mg/mL) 0.20 mL 50% Dextran Sulfate 0.02 mL 50x Denhardt’s
OR
1. Alternative prehybridization buffer (4xSSC ; 50% deionized formamide) ; Example. we need 10 ml prehyb.buffer deionized formamide = 5 ml 20xSSC(4xSSC) = 2 ml Add DEPC H2O = 3 ml
2. Fill slide case with buffer and slides and incubate w/o coverslips 1 hr at 370 C Or 2 hr at RT in humid chamber.(Alternative incubate at least 10 min)

Hybridization

~ 30 min.
1. Set oven to 53 *C. Discard prehybridization buffer and rinse slides in 2x SSC and briefly dry prior to hybridization.
2. Add probe to prehyb. buffer at a concentration of 1:373
3. Add probe to tissue sections. [300 ul each]
*4. Denature tissue sections with probe for 10 min at 100 oC [*with a slide griddle attached to a PTC-100 thermal controller]. Use hot plate for our lab.(Alternative)
5. Cover tissue sections with Dnase-free coverslips.
6. Incubate slides at 53 oC O/N in a humid chamber overnight.
7. Place 2x SSC, 1x SSC, and 0.5x SSC in their containers and heat them in a 40 *C oven overnight.

Day 2

Carefully remove cover slips from sections by immersing slides for 5-10 min in 2xSSC at RT
1. Wash slides in 2x SSC twice for 15 min each at 40 oC.
2. Wash in 1x SSC three times for 15 min at 40 oC.
3. Wash in 0.5 x SSC for 15 min at 40 oC
4. Equilibrate tissues in Buffer 1 (100 mM tris-HCl, 10 mM NaCl, pH 7.5) for 10 min
5. Block tissues with Blocking Buffer (Buffer 1 + 2% sheep serum + 0.3% Triton X- 100) for 1 hr at RT. {500ul each}


Detection

1. Prepare Alkaline phosphatase (AP)-labeled sheep anti-DIG antibody conjugate. Dilute antibody 1:1000 in Buffer 1 + 1% Sheep serum + 0.3% Triton X-100.
2. Add solution (0.5 -1 mL) to dry section and incubate without coverslip at RT For 2 hours in a humid chamber.
3. Rinse slides with Buffer 1 for 2 x 10 min.
4. Rinse slides in Buffer 2 (100 mM tris-HCl, 100 mM NaCl, 50 mM MgCl2, pH 9.5 – make sure!!) for 10 min.
5. Prepare substrate by adding 45 ul NBT (nitroblue tetrazolium) + 35 ul BCIP to 10 mL of Buffer2
6. Incubate tissue sections with substrate solution 30 min -1 hour at RT in humid chamber. Check tissues after 15 min. to see if color has shown up. If color does not show up at the end of an hour, incubate overnight
7. Rinse slides in 3x dH2O.
8. Counter stain slides with 0.05% aqueous Bismarck Brown Y for 3 min.
9. Rinse slides with dH2O, then 70% and 100% EtOH breifly.
10.Let slides air dry then permount.

Note: To optimize the protocol the following variables can be modified:
1. Concentration and length of solubilization with proteinase K.
2. Dilution of probe and anti-DIG-AP antibody conjugate
3. Temperature and length of incubation of anti-DIG antibody
4. Temperature and stringency of washes
5. Length of incubation of BCIP-NBT substrate. Signals should develop beginning at 30 min to 1 h. Do not incubate beyond 1 h.

Reagents For In-situ:

I. Abalone Rickettsia Okigonucleotide Primers

Optimum oligonucleotide conc. = 200 pm/ul
Primer = Conc (pmole/ul)
RA 5-1 = 234

Have 234 pmole/ul (X) = want 200 pmole/ul x 200 ul?
X = 170.94 ul

RA 3-6 183 = 218.57 ul
RA 3-8 151 = 264.90 ul
RA 5-6 238 = 168.067 ul

Total = 822.477 ul

Then:
RA 5-1: Have 822.477ul (X) = want 0.992 ul x 170.94ul
X = 0.206 ul

RA 3-6: = 0.264 ul

RA 3-8 = 0.319 ul

RA 5-6 = 0.203 ul

Total = 0.992ul

II. Control group

= 5 ul viral # 6 instead of the primers
4 ul Dnase free-water
Then add reagents (11 ul).
So the total of the Labeling procedure is 20 ul.

III. Permeabilization of tissues

Wash tissues with 50 ug/ml of Proteinase K in Tris-buffer for 45 min at 37 C.
Working 50 ug/ml of Proteinase K in Tris-buffer.

Proteinase K: stock 20 mg/ml = 20,000 ug/ml

Example: Have 20 mg/ml (20,000 ug/ml) (X) want 50 ug/ml x 2,000 ul
X = 50 ug/ml x 2,000 ul/20,000 ug/ml
= 0.005 ml
Take Protein K = 5 ul
Tris buffer = 2,000 – 5 = 1,995 ul

We Use about 15 ml/4 slides

Then we want 15 ml of 50 ug/ml Proteinase- K in Tris-buffer
Have 20,000 ug/ml (X) want 50 ug/ml 15 ml
X = 50 ug/ml x 15,000ul /20,000 ug/ml
Take Protein K from stock = 37.5 ul
Tris-buffer =15,000 ul -37.5 ul = 14.97 ml

IV. Hybridization

Probe : Prehybridization buffer = 1:373 (Dolly recommend: 13:4850 )

Keeping a Laboratory NotebookThe most important piece of equipment you will use when working in lab is your lab notebook. Since science is built upon the premise that results are reproducible, we must leave detailed information so others could reproduce our work if they read our notes. Your notebook is the only source of information of all that you have done in lab. It should be an accurate account of what you did, why you did it, when you did it, what the results were, and what these results mean.
Here are a few simple rules to keep in mind while developing your notebook:
1. Notebooks should always be legible. Record the date at the top of each page. Also, a notebook is not designed to be a duplication of the lab manual. Just cite the lab manual pages, and then record any deviations from the published protocol.
2. Use ONLY dark blue or black pen. Some highlights and colored pen is OK for emphasizing some parts.
3. Record any measurements or calculations you perform.
4. Explain why your are performing a particular procedure. What is the purpose and what do you expect to see? Is there more than one possible outcome?
5. Record the data. If it is a gel, then tape a copy of the gel into your notebook, label the lanes (by numbers), and link the gel photo to the appropriate date. For example, if you did a PCR in week 2 and run the gel in week 3, then indicate when the samples where prepared near the gel. Label the percentage of the gel and what was loaded in each lane (i.e. tell what the numbers for each lane mean).
6. It is very important to record your interpretations of the results. What do all those little white lines mean??!
7. You should note if you need to repeat the experiment, or you are ready for the next step. At this time, you should mention what the next step is.
8. Record any observations that you think might be significant. If you deviated from the normal protocol, you should record all variations.
9. It is often a good idea to record the times when you start and stop a procedure. This will enable you to be sure you do not go too long or too short.
10. It is essential to keep complete records of ALL experiments, whether "successful" or "unsuccessful". It is amazing how often it happens that an experiment initially judged "unsuccessful" later proves to have been "successful" in an unanticipated way.
11. If you have an interesting idea about an experiment you've already done or about an experiment that you could do, write it down in the notebook immediately, after entering the date.
12. If someone shows you how to operate an instrument or carry out a procedure that is new to you, take notes in your notebook. You can use these notes when you later go back to do the procedure on your own.
13. Your primary guiding principle in the laboratory should be to maintain your notebook so carefully and completely that, at some later time, you or any other scientist could repeat any experiment or operation using only your lab notebook as a resource.

Mini Prep: Alkaline Lysis Protocol

(R. Estes, 12/03)
1. Spin down your cells. Your DNA is still in the cells, so it is in the pellet at this stage.
2. Discard the supernatant. Pieces of cell wall are released from the bacteria and are floating around in the supernatant. These cell wall pieces can inhibit enzyme action on your final DNA, so it is important to get rid of all of the supernatant and to even invert the tube and wipe the lip with a Kim-wipe or Q-tip.
3. Resuspend the cells in buffer (often Tris) and EDTA. EDTA chelates divalent metals (primarily magnesium and calcium). Removal of these cations destabilizes the cell membrane. It also inhibits DNases. Glucose should also be added to maintain osmolarity and prevent the buffer from bursting the cells.
4. Lyse the cells with sodium hydroxide (NaOH) and SDS. This highly alkaline solution gave rise to the name of this technique. Mix this by gentle inversion and incubate on ice for five minutes (but no longer, or your DNA will be irreversibly denatured). Three things happen during this stage:
a. SDS pops holes in the cell membranes. SDS (sodium dodecyl (lauryl) sulfate) is a detergent found in many common items such as soap, shampoo and toothpaste.
b. NaOH loosens the cell walls and releases the plasmid DNA and sheared cellular DNA.
c. NaOH denatures the DNA. Cellular DNA becomes linearized and the strands are separated. Plasmid DNA is circular and remains topologically constrained.
5. Renature the plasmid DNA and get rid of the garbage. Add potassium acetate (KAc), which does three things:
a. Circular DNA is allowed to renature. Sheared cellular DNA remains denatured as single stranded DNA (ssDNA).
b. The ssDNA is precipitated, since large ssDNA molecules are insoluble in high salt.
c. Adding sodium acetate to the SDS forms KDS, which is insoluble. This will allow for the easy removal of the SDS from your plasmid DNA.
Now that you've made it easy to separate many of the contaminants, centrifuge to remove cell debris, KDS and cellular ssDNA. Your plasmid DNA is in the supernatant, while all of the garbage is in the pellet.
6. Precipitate the plasmid DNA by alcohol precipitation (ethanol or isopropanol) and a salt (such as ammonium acetate, lithium chloride, sodium chloride or sodium acetate) and spin this down. DNA is negatively charged, so adding a salt masks the charges and allows DNA to precipitate. This will place your DNA in the pellet.
7. Rinse the pellet—your plasmid DNA—in ice-cold 70% EtOH and air-dry for about 10 minutes to allow the EtOH to evaporate.
8. Resuspend your now clean DNA pellet in buffer (often Tris) and EDTA plus RNases to cleave any remaining RNA. Your DNA is now back in solution.
DNA of this purity is good for a number of uses, such as in vitro transcription or translation or cutting with some enzymes. If you are sequencing or transforming this DNA into mammalian cells, you'll want to use additional purification techniques such as phenol extraction, Qiagen column purification, or silica-based purification.

Mini-Prep: Alkaline Lysis Method
(R. Estes, 12/03)
Alkaline lysis is the method of choice for isolating circular plasmid DNA, or even RNA, from bacterial cells. It is probably one of the most generally useful techniques as is a fast, reliable and relatively clean way to obtain DNA from cells. If necessary, DNA from an alkaline lysis prep can be further purified.
Alkaline lysis depends on a unique property of plasmid DNA. It is able to rapidly anneal following denaturation. This is what allows the plasmid DNA to be separated from the bacterial chromosome.
Typically, you will grow up E coli cells that contain the plasmid you want to isolate, then you will lyse the cells with alkali and extract the plasmid DNA. The cell debris is precipitated using SDS and potassium acetate. This is spun down, and the pellet is removed. Isopropanol is then used to precipitate the DNA from the supernatant, the supernatant is removed, and the DNA is resuspended in buffer (often TE). A mini prep usually yields 5-10 ug. This can be scaled up to a midi prep or a maxi prep, which will yield much larger amounts of DNA (or RNA).
Specific protocols for alkaline lysis differ widely from lab to lab, and even from scientist to scientist. The basic principles behind the procedure, however, are fairly uniform.

Nitzchia Culture


To start a working culture from bacto-agar stock: Add 50ml of bacto-agar stock to 500ml of seawater nutrient. (A tenth of the receiving volume is recommended to start a working culture. However, adjusting the amount of inoculant at this point will determine the time before the working culture is ready to be used - less will prolong the culture from maturing and more will shorten the ripening period). Make sure to store working culture covered in 15°C incubator with constant light and an airstone. Label flask with contents, date, and name.

Water changes: Approximately every 3 days 50ml of ripe working culture should be added to 500ml of fresh seawater nutrient. Label new flask with contents, date, and name every time a water change is made.

OTC Assay Protocol

1. Preparation of Plates

1.1. Culture of Bacillus cereus.
1.1.1. Materials.
(1) .15g BD BactoTryptic Soy Broth
1.1.2. Protocol.
(1) Prepare 5mL TSB media
(2) Inoculate by touching sterile pipet tip in B.cereus stock and placing into media.
(3) Incubate at room temperature,rotating at 120rpm, overnight.
1.2. Preparation of Mueller Hinton Agar.
1.2.1. Materials.
(1) 17.1g BDBBL Mueller Hinton II agar
1.2.2. Protocol.
(1) Add 17.1g Mueller Hinton agar to 450mL NanoPureH20and swirl to mix.
(2) Autoclave on appropriate liquid cycle.
(3) Place in 48C water bath for approximately 1 hour.
1.3. Inoculation of B. cereus.
1.3.1. Materials.
(1) 1100×15mm petri dish per sample
1.3.2. Protocol.
(1) Add 4.05mL B.cereus suspension to cooled (48C) Mueller Hinton agar and swirl to mix.
(2) Add 10mL of inoculated agar to each labelled plate, careful to avoid bubbles.
(3) Swirl agar to form uniform layer and allow to solidify.

2. Preparation of samples and standards

2.1. Preparation of Tissue and/or Feed Homogenate.
2.1.1. Materials.
(1) Tissue samples (foot and/or digestive gland)
(2) 7mLTenbroeckTissue Grinder, KONTES
(3) 13.6g Sigma P-5379 Potassium Phosphate
(4) 0.22µm filter
(5) 2-15mL conical tubes per sample
2.1.2. Protocol.
(1) Prepare 0.1M KH2PO4
(2) Add approximately 0.25g tissue (prepare foot samples by dicing and smashing with rubber mallet; prepare feed samples by smashing with rubber mallet) to homogenizer and suspend in KH2PO4 buffer (16mL to 1g tissue).
(3) Homogenize by twisting pestle until sample is completely homogenized (no visible tissue or particles).
(4) Add homogenate to fresh, labeled 15mL conical tube.
(5) Centrifuge tubes at approximately 2000rpm for at least 10 minutes; repeat if separation does not appear complete.
(6) Decant supernatant into fresh, labeled 15mL conical tube.
(7) Store at 80°Cif necessary.
2.2. Preparation of OTC Standards.
2.2.1. Materials.
(1) 0.01g Sigma O-5750 Oxytetracycline Dihydrate
(2) 20mL 0.1N HCl
(3) 0.22µm filter
(4) 0.1M KH2PO4 buffer
2.2.2. Protocol.
Prepare500ppm stock solution by dissolving 0.01g OTC Dihydrate in 20mL 0.1N HCl and filter sterilizing (may be stored at 4°C for up to three days or at80°C indefinitely).
(2)Prepare 50ppm stock by adding 500ppm stock to 0.1NHClin ratio of 1mL stock to 9mL HCl in labelled tube.
(3)Prepare 5ppm stock by adding 50ppm stock to KH2PO4buffer in ratio of 1mL stock to 9mL buffer.
(4) Prepare 20mL 0.2ppm stock for in-plate standards by adding 80µLof 50ppm stock to 19.92mL KH2PO4 buffer.
(5)Prepare standards from 0–1 ppm by adding 5ppm stock in following amounts
Concentration (ppm)
µL5ppm stock
µLKH2PO4 buffer
0.0
0.1
0.3
0.5
0.7
0.9
1.0
0
20
60
100
140
180
200
1000
980
940
900
860
820
800

3. Adding samples and standards to plate

3.1. Materials.
(1) Forceps
(2) 48mm × 10mm penicylinders per plate
(3) Tissue homogenates
(4) OTC standards
3.2. Protocol.
(1) Using sterile forceps place 1 penicylinder on top of cooled agar in each quarter of plate.
(2) Lightly tap the cylinder into the agar, placing the bottom just below the agar’s surface.
(3) Add 250µL of 0.2ppm standard to1 cylinder of each plate, carefully noting which has standard.
(4) Add 250µLof sample homogenate or standard to each of the remaining3cylinders.
(5) Cover and incubate plates for approximately 20 hours.

4. Measuring zones of inhibition

4.1. Materials.
(1) Forceps
(2) Vernier calipers
4.2. Protocol.
(1) Using forceps, remove cylinders from plate.
(2) Turn plate upside down and place on dark surface (e.g. lab bench).
(3) Use calipers to measure diameter of zone, measured along radius of plate itself.

5. Quantifying OTC concentrations

5.1. Protocol.
(1) Enter zone diameters for sample and 0.2ppm standard for each plate.
(2) Calculate mean diameter of zones for each sample.
(3) Calculate ‘grand’ mean of 0.2ppm standard for all samples.
(4) Calculate multiplier for each sample by the equation
Multiplier = 0.2ppm grand mean
0.2ppm sample diameter
(5) Plot diameter vs. concentration of known OTC standards.
(6) Add linear regression of standard curve, setting intercept at mean diameter of 0ppm zones.
(7) Calculate OTC concentration according to standard curve, multiplying to account for any dilution used.

PCR Protocol for Withering Syndrome (RLP)

(R. Estes, 9/04)
1. Wipe down workbench with 10% bleach then wipe off with deionized water to remove bleach. Let dry overnight.
2. Label sample tubes. Use 0.2ml thin-walled PCR tubes.
3. Thaw all components of cocktail (except Taq) at room temperature.
4. Aliquot approximately 100 ng of template into PCR tube. If the concentration template is 50 ng/ml, use 2 ml.
5. Preheat the thermal cycler to 95°C.
6. Prepare cocktail of PCR components. Include a positive and negative template control.

Stock
End
Per
Reagent
Concentration
Concentration
Tube (ml)
5 X Buffer
5x
1
X
4
MgCl2
25 mM
1.5
mM
1.2
BSA
10 mg/ml
400
ng/ml
0.8
H2O



11.08
dNTP's
10 mM
200
mM
0.4
RA 3-6
100 pmol/ml
0.5
mM
0.1
RA 5-1
100 pmol/ml
0.5
mM
0.1
Taq*
5 U/ml
1.6
U
0.32
Template



2 (100 ng)
Total Reaction Volume
20 ml
* Promega Taq Polymerase in Storage Buffer A
7. Once the Taq is added to the cocktail immediately dispense cocktail into the tubes containing template. Using a separate pipette tip for each tube and gently mixing ( by pipetting up and down) the cocktail mixture between aliquots.
8. Place tubes in preheated (95°C) thermal cycler, then start program.
Thermal Cycler Program = RLP4

Time
Temp (°C)
Step 1
3 min
95
Step 2
1 min
95
Step 3
30 sec
62
Step 4
30 sec
72
Repeat steps 2-4, 40 times
Step 5
10 min
72

PCR Amplification of Genomic DNA

Note: Protocol using Promega Taq DNA Polymerase #M1861
Thermocycler used is MJ Research MiniCycler.
MgCl2 concentration determined 3/08/2000, JLM

1) Thaw all components of cocktail (except Taq) at room temperature.
2) Once thawed place cocktail components on ice.
3) Label sample tubes. Use 0.2ml thin-walled eppendorph tubes.
4) Aliquot appropriate amount of template into a 0.2ml thin-walled eppendorph tube. For genomic template use 8ng/ul. For InstaGene extractions use 10.0ul per 100ul PCR reaction volume. Place the sample tubes on ice.
5) Preheat the thermocycler to 95C.
6) Prepare cocktails of PCR components as described below.
Prepare cocktails omitting template (DNA).
Prepare enough cocktail for all sample tubes plus 10% extra. Include a positive and negative template control. Note all lot numbers of the cocktail components in your lab notebook.
Volumes below are per tube.
Keep the cocktail on ice.
Prepare in the following order while on ice:
10.0ul 5.0ul 10X rxn buffer (Promega MgCl2 free buffer)
8.0ul 4.0ul 25mM MgCl2 (Promega)
10.0ul 5.0ul 50% Acetamide
Adjust accordingly PCR H2O
Gently mix contents via pipetting up and down, avoiding air bubbles.
0.8ul 0.4ul 25mM dNTP Mix (25mM/ea)
1.0ul 0.50ul 50uM Forward Primer (8fm or 8fm-HEX)
1.0ul 0.50ul 50uM Reverse Primer (1492R-rph)
Gently mix contents via pipetting up and down, avoiding air bubbles.
1.2ul 0.6ul Taq DNA Polymerase (Promega)
100.0ul 50.0ul Total reaction volume per tube (including template/cocktail)
7) Immediately dispense cocktail into the tubes containing template. Using a separate pipette tip for each tube and gently mixing the cocktail mixture between aliquots.
8) Add two drops of mineral oil to each tube.
9) Run reactions immediately, keeping them on ice while transporting.
Thermocycler Program: 16S-JLM
1) 94C 00:03:00
2) 94C 00:01:00
3) 50C 00:01:30
4) 72C 00:02:00 +5 sec/cycle
5) to step 2, 28 times
6) 94C 00:01:00
7) 50C 00:01:30
8) 72C 00:07:00
9) 4C 00:00:00
10) END

PCR Amplification of Plasmid DNA

(“James Gray 2/98” modified 4/15/99 JLM)
Note: Protocol using Promega Taq DNA Polymerase #M1861
Thermocycler used is MJ Research MiniCycler.
MgCl2 concentration determined 3/10/98, JLM
1) Thaw all components of cocktail (except Taq) at room temperature.
2) Once thawed place cocktail components on ice.
3) Label sample tubes. Use 0.2ml thin-walled eppendorph tubes.
4) Aliquot appropriate amount of template into a 0.2ml thin-walled eppendorph tube. For plasmid template use 4.0pg/ul. Place the sample tubes on ice.
5) Preheat the thermocycler to 95C.
6) Prepare cocktails of PCR components as described below.
Prepare cocktails omitting template (DNA).
Prepare enough cocktail for all sample tubes plus 10% extra. Include a positive and negative template control. Note all lot numbers of the cocktail components in your lab notebook.
Volumes below are per tube.
Keep the cocktail on ice.
Prepare in the following order while on ice:
10.0ul 5.0ul 10X rxn buffer (Promega MgCl2 free buffer)
8.0ul 4.0ul 25mM MgCl2 (Promega)
10.0ul 5.0ul 50% Acetamide
Adjust accordingly PCR H2O
Gently mix contents via pipetting up and down, avoiding air bubbles.
0.8ul 0.4ul 25mM dNTP Mix (25mM/ea)
1.0ul 0.50ul 50uM Forward Primer
1.0ul 0.50ul 50uM Reverse Primer
Gently mix contents via pipetting up and down, avoiding air bubbles.
1.2ul 0.6ul Taq DNA Polymerase (Promega)
100.0ul 50.0ul Total reaction volume per tube (including template/cocktail)
7) Immediately dispense cocktail into the tubes containing template. Using a separate pipette tip for each tube and gently mixing the cocktail mixture between aliquots.
8) Add two drops of mineral oil to each tube.
9) Run reactions immediately, keeping them on ice while transporting.
Thermocycler Program: 16S-JLM
1) 94C 00:03:00
2) 94C 00:01:00
3) 50C 00:01:30
4) 72C 00:02:00 +5 sec/cycle
5) to step 2, 28 times
6) 94C 00:01:00
7) 50C 00:01:30
8) 72C 00:07:00
9) 4C 00:00:00
10) END

PCR Primer Design

(R. Estes, 12/03)
The sequences of the primers used for PCR amplification will have a major effect on the specificity and sensitivity of the reaction. When designing two PCR amplification primers, the following guidelines should be considered:
· Primers should be typically 20-30 bases long.
· Primers should not contain bases complementary to themselves or with each other, especially at the 3' and 5' ends. When this occurs it can promote for formation of PCR artifacts called primer-dimers.
· The melting temperatures (Tm) should be 50-70 C°, so that high annealing temperatures can be used.
· Primer pairs with similar Tm should be created.
· A 40% - 60% G+C content is often recommended for both primers; for primers with a G+C content much less than 40%, the primer sequence might need to be extended beyond 20 bases to keep the melting temperature above the recommended lower limit of 50 C°.
· Avoid long runs of a single base, as this may cause secondary hybridization on targets that happen to contain the complementary motif.
· The 3' end should contain a G or C to stabilize this end.
· Avoid designing primers that have secondary structures.
· Use primer design computer programs when designing custom primers, they are good predictors of internal relationships. Try the program at the following website to check the quality of your primers.
http://www.premierbiosoft.com/netprimer/netprlaunch/netprlaunch.html

General PCR Protocol

(R. Estes, 12/03)
1. Bleach pipettes by wiping with moist cloth with 10% bleach and then wipe off with deionized water to remove bleach. Dry overnight before using. [Do not pour/spray/drip bleach directly onto pipets. This can harm the pipets.]
2. Wipe down workbench with 10% bleach then wipe off with deionized water to remove bleach. Let dry overnight. [Do not use a spray bottle as the bleach gets everywhere.]
3. Put on lab coat and gloves.
4. Wipe down workbenck with 70% EtOH to remove dirt and dust. Let dry for 10 minutes.
5. Thaw the following reagents: MgCl2 [OK to briefly vortex reagents]
10X buffer
BSA
dNTPs
Primers (A3 and A4) [no need to vortex]
Thaw DNA sample templates. [Pipet up and down to mix]
Leave Taq in freezer until needed. [Do not mix]
6. Add proper amount of reagents into a 1.5mL tube using filter tips. [Always use new pipet tip between reagents and/or samples.]
7. Add taq to tube. Gently pipet up and down until you see glycerol mixed into solution. [Do not vortex]

8. Add 24µL of mastermix into each PCR tube.
9. Add 1µL of DNA template into each of the PCR tubes.
10. If needed, it is ok to do a quick spin of the tubes. [This means you get the centrifuge up to speed (1500 RPM on large centrifuge) and let it go back down right away]
11. Heat up thermal cycler to 95°C.
12. Place plate in thermal cycler [preheated to to 95°C].
13. Run thermal cycler program for HERPES.
14. For PCR 2, use the same procedure as above with primers A5 and A6 and add 1 ml of product from PCR 1.
15. Store samples in refrigerator until ready to run the gel.

General PCR Protocol for Oyster Herpes Virus

(R. Estes, 12/03)
1. Bleach pipettes by wiping with moist cloth with 10% bleach and then wipe off with deionized water to remove bleach. Dry overnight before using. [Do not pour/spray/drip bleach directly onto pipets. This can harm the pipets.]
2. Wipe down workbench with 10% bleach then wipe off with deionized water to remove bleach. Let dry overnight. [Do not use a spray bottle as the bleach gets everywhere.]
3. Put on lab coat and gloves.
4. Wipe down workbenck with 70% EtOH to remove dirt and dust. Let dry for 10 minutes.
5. Thaw the following reagents: MgCl2 [OK to briefly vortex reagents]
10X buffer
BSA
dNTPs
Primers (A3 and A4) [no need to vortex]
Thaw DNA sample templates. [Pipet up and down to mix]
Leave Taq in freezer until needed. [Do not mix]
6. Add proper amount of reagents into a 1.5mL tube using filter tips. [Always use new pipet tip between reagents and/or samples.]
7. Add taq to tube. Gently pipet up and down until you see glycerol mixed into solution. [Do not vortex]

8. Add 24µL of mastermix into each PCR tube.
9. Add 1µL of DNA template into each of the PCR tubes.
10. If needed, it is ok to do a quick spin of the tubes. [This means you get the centrifuge up to speed (1500 RPM on large centrifuge) and let it go back down right away]
11. Heat up thermal cycler to 95°C.
12. Place plate in thermal cycler [preheated to to 95°C].
13. Run thermal cycler program for HERPES.
14. For PCR 2, use the same procedure as above with primers A5 and A6 and add 1 ml of product from PCR 1.
15. Store samples in refrigerator until ready to run the gel.

Making Plasmid

(Last updated 8-12-08)
When testing multiple sets of primers make sure your plasmid will contain all sets, include the largest range of your amplified product.
1. Run a conventional PCR on an extracted sample using your primers. Visualize your product on a gel to make sure you have clean, distinct bands (don’t forget a negative control)! If all looks good, you will use this product for cloning your plasmid.
2. Before cloning your plasmid you need to clean up/purify your PCR product using the Qiaquick kit (Qiagen) following the protocol on page 18 of the kit manual.
3. After purifying your product, quantify the amount of DNA in your sample using the microplate reader. For this you should follow the DNA quanting via standards in layout protocol (ask for this). Make sure you quant your purified product in duplicate or triplicate and take the average of all (if your r-value and standards are acceptable). This is important because you want to know your DNA concentration prior to cloning.
4. At this point you may want to send your purified product out for sequencing. See Basic Protocol for direct DNA sequencing. When you receive your sequence back, blast it against the NCBI database to make sure you are amplifying the correct target.
5. Make sure you have several LB plates with 50 µg/ml ampicillin already prepared for culturing your clones. Check for extra plates in the fridge. Do not use plates that are more than 6 weeks old. If you need to prepare fresh plates the cocktail for LB media is in the recipe book.
6. You are now ready to clone using the TOPO TA Cloning Kit for Sequencing (Invitrogen). The kit is stored in the -20°C freezer. You will need a cooler with ice, a 42°C water/dry bath, and a 37°C incubator with a shaking platform for part B of the cloning process.
A. Performing the TOPO Cloning Reaction (pg 5 of user manual): We will use half reactions for chemically competent E. coli
· Add 2µl purified PCR product, 0.5µl salt solution, and 0.5µl TOPO vector into a PCR tube – Make sure to load reagents in this order!!
· Mix gently by flicking tube and giving it a brief spin down
· Incubate reaction at room temp for 10 min
· After 10 min, place the reaction on ice.
(NOTE: You may store the TOPO Cloning reaction at -20°C overnight)
B. Transforming with One Shot TOP10 E. coli (pg 9 of user manual):
· Thaw on ice 1 vial of Top10 E.coli cells (stored in -80°C freezer) for 5 min.
· Add 1µl of TOPO cloning reaction (from part A) to ½ vial (25µl) of the E. coli cells. (Refreeze remaining E. coli cells in -80°C and freeze remaining cloning reaction in -20°C)
· Mix gently by flicking tube and giving it a brief spin down
· Incubate on ice for 15 min
· Heat shock cells @ 42°C for 30 sec
· IMMEDIATELY place cells back on ice for 2 min
· Add 125µl of room temp SOC medium (check chemical shelf in 240)
· Cap tube tightly and incubate @ 37°C on shaking platform for 1 h
· Warm LB-amp plates for 30 min @ 37°C
· After the cells have incubated for an hour, spread 30-50µl of product on prewarmed LB-amp plates. Plate 2 or 3 different volumes to ensure at least one plate will have well spaced colonies.
· Incubate plates overnight @ 37°C
C. Preparing Single Colony Clones:
· Prewarm more LB-amp culture plates @ 37°C for 30 min for growing single colonies (patch plate)
· Prepare a 15ml falcon tube with 5ml of LB liquid media and 25µl of 200X stock ampicillin in duplicate. You will need to make 2 tubes for every single colony you plan to culture (one to purify and one for long term storage) plus 1 tube as a blank control.
· Take your plates you incubated overnight which now should have lots of colonies on them and using a pipette tip pick only one colony off of each plate, touch the tip to the fresh patch plate, then dip the tip into the appropriate falcon tube with liquid LB-amp media.
· Grow these single colony cultures overnight at 37°C making sure the falcon tubes are on a shaking platform.
· Parafilm and store multiple colony plates in the fridge (4°C)

D. Analyzing Transformants by PCR: (pg 13 of user manual)
· Prepare your master mix following your original PCR recipe except substitute 2µl of water in place of your product.
· Aliquot 25µl of your master mix into PCR tubes. Make enough reactions for each colony and a negative control.
· Take your single colony culture and using a pipette tip touch your colony and dip the tip into your PCR tube.
· Run your PCR on the thermocycler and then visualize on a gel to make sure your product is the correct size.
· Parafilm single colony LB-amp plates and store in the fridge (4°C)

E. Long Term Storage (pg 13 of user manual):
· Add 880µl of glycerol to 1 set of cultures in your falcon tubes
· Store at -80°C

F. Purifying your Plasmid
· Centrifuge the remaining falcon tube for 10 min @ 2500rpms @ 4°C to pellet cells.
· Pour off LB-amp media
· And finally….follow protocol on pg 22 of the Qiaprep Spin Mini Prep Kit to purify your plasmid!

CONGRATULATIONS! YOU ARE NOW A PLASMID MASTERMAKER! :o)


Restriction Enzyme Overview

(New England Biolabs, Edited by R.E. Strenge 6/04)
Restriction enzymes were discovered about 30 years ago during investigations into the phenomenon of host-specific restriction and modification of bacterial viruses. Bacteria initially resist infections by new viruses, and this "restriction" of viral growth stemmed from endonucleases within the cells that destroy foreign DNA molecules. Among the first of these "restriction enzymes" to be purified were EcoR I and EcoR II from Escherichia coli, and Hind II and Hind III from Haemophilus influenzae. These enzymes were found to cleave DNA at specific sites, generating discrete, gene-size fragments that could be re-joined in the laboratory. Researchers were quick to recognize that restriction enzymes provided them with a remarkable new tool for investigating gene organization, function and expression.
As the use of restriction enzymes spread among molecular biologists in the late 1970’s, companies such as New England Biolabs began to search for more. Except for certain viruses, restriction enzymes were found only within prokaryotes. Many thousands of bacteria and archae have now been screened for their presence. Analysis of sequenced prokaryotic genomes indicates that they are common--all free-living bacteria and archaea appear to code for them.
Restriction enzymes protect bacteria from infections by viruses, and it is generally accepted that this is their role in nature. They function as microbial immune systems. When a strain of E.coli lacking a restriction enzyme is infected with a virus, most virus particles can initiate a successful infection. When the same strain contains a restriction enzyme, however, the probability of successful infection plummets. The presence of additional enzymes has a multiplicative effect; a cell with four or five independent restriction enzymes could be virtually impregnable.
Restriction enzymes usually occur in combination with one or two modification enzymes (DNA-methyltransferases) that protect the cell’s own DNA from cleavage by the restriction enzyme. Modification enzymes recognize the same DNA sequence as the restriction enzyme that they accompany, but instead of cleaving the sequence, they methylate one of the bases in each of the DNA strands. The methyl groups protrude into the major groove of DNA at the binding site and prevent the restriction enzyme from acting upon it. Together, a restriction enzyme and its "cognate" modification enzyme(s) form a restriction-modification (R-M) system. In some R-M systems the restriction enzyme and the modification enzyme(s) are separate proteins that act independently of each other. In other systems, the two activities occur as separate subunits, or as separate domains, of a larger, combined, restriction-and-modification enzyme.
Restriction enzymes are traditionally classified into three types on the basis of subunit composition, cleavage position, sequence-specificity and cofactor-requirements. However, amino acid sequencing has uncovered extraordinary variety among restriction enzymes and revealed that at the molecular level there are many more than three different kinds.
Type IType I enzymes are complex, multisubunit, combination restriction-and-modification enzymes that cut DNA at random far from their recognition sequences. Originally thought to be rare, we now know from the analysis of sequenced genomes that they are common. Type I enzymes are of considerable biochemical interest but they have little practical value since they do not produce discrete restriction fragments or distinct gel-banding patterns.
Type IIType II cut DNA at defined positions close to or within their recognition sequences. They produce discrete restriction fragments and distinct gel banding patterns, and they are the only class used in the laboratory for DNA analysis and gene cloning. Rather then forming a single family of related proteins, type II enzymes are a collection of unrelated proteins of many different sorts. Type II enzymes frequently differ so utterly in amino acid sequence from one another, and indeed from every other known protein, that they likely arose independently in the course of evolution rather than diverging from common ancestors.
The most common type II enzymes are those like Hha I, Hind III and Not I that cleave DNA within their recognition sequences. Enzymes of this kind are the principle ones available commercially. Most recognize DNA sequences that are symmetric because they bind to DNA as homodimers, but a few, (e.g., BbvC I: CCTCAGC) recognize asymmetric DNA sequences because they bind as heterodimers. Some enzymes recognize continuous sequences (e.g., EcoR I: GAATTC) in which the two half-sites of the recognition sequence are adjacent, while others recognize discontinuous sequences (e.g., Bgl I: GCCNNNNNGGC) in which the half-sites are separated. Cleavage leaves a 3´-hydroxyl on one side of each cut and a 5´-phosphate on the other. They require only magnesium for activity and the corresponding modification enzymes require only S-adenosylmethionine. They tend to be small, with subunits in the 200–350 amino acid range.
The next most common type II enzymes, usually referred to as ‘type IIs" are those like Fok I and Alw I that cleave outside of their recognition sequence to one side. These enzymes are intermediate in size, 400–650 amino acids in length, and they recognize sequences that are continuous and asymmetric. They comprise two distinct domains, one for DNA binding, the other for DNA cleavage. They are thought to bind to DNA as monomers for the most part, but to cleave DNA cooperatively, through dimerization of the cleavage domains of adjacent enzyme molecules. For this reason, some type IIs enzymes are much more active on DNA molecules that contain multiple recognition sites.
The third major kind of type II enzyme, more properly referred to as "type IV" are large, combination restriction-and-modification enzymes, 850–1250 amino acids in length, in which the two enzymatic activities reside in the same protein chain. These enzymes cleave outside of their recognition sequences; those that recognize continuous sequences (e.g., Eco57 I: CTGAAG) cleave on just one side; those that recognize discontinuous sequences (e.g., Bcg I: CGANNNNNNTGC) cleave on both sides releasing a small fragment containing the recognition sequence. The amino acid sequences of these enzymes are varied but their organization are consistent. They comprise an N-terminal DNA-cleavage domain joined to a DNA-modification domain and one or two DNA sequence-specificity domains forming the C-terminus, or present as a separate subunit. When these enzymes bind to their substrates, they switch into either restriction mode to cleave the DNA, or modification mode to methylate it.
Type IIIType III enzymes are also large combination restriction-and-modification enzymes. They cleave outside of their recognition sequences and require two such sequences in opposite orientations within the same DNA molecule to accomplish cleavage; they rarely give complete digests. No laboratory uses have been devised for them, and none are available commercially.

Western Blot

Components of Western Blot:
1) Sample Preparation
2) Polyacrylamide gel and transfer to nitrocellulose membrane
3) Probe membrane with antibodies
4) photograph and score
Step one: Polyacrylamide Gel casting (Resolving gel and stacking gel)
Cast gel using OWL casting device
Helpful hints:
· Clean all surfaces with 70% ethanol (unclean surfaces cause air bubbles)
· Make sure that the gel caster is level
· Place spacers between glass plates, with the glass plates in the plastic bags provided
· Assemble gel caster such that the order is: plastic plate, bag with glass plates and spacers, plastic plate, thick acrylic spacer.
· Tighten gel caster knobs firmly; but not too firmly—the glass plates are breakable
· Be sure that separators and combs are the same thickness (1.5 mm)
1) Mix Resolving gel according to attached recipe.
Helpful Hints:
· Make fresh ammonium persulfate EVERY time
· Heat SDS for 15 seconds to liquify
· Add temed last and under the hood (it’s stinky)
o Temed is the ingredient that causes rapid polymerization
2) Pour gel
· I used 1000 uL pipetter. Be careful not to allow any bubbles to form. You have approximately 5 minutes to pour the gel before polymerization occurs.
3) Overlay gel with Butanol
· Don’t forget this step. This creates a smooth line between gel layers.
4) Let polymerize for 1.5 hours
· Leave seated on the bench
Stacking Gel:

1) Pour off Butanol and rinse gel 3 times with dH2O
2) Mix stacking gel according to attached recipe
3) Pour gel with 1000 uL pipetter
4) Place 1.5 mm combs into the top of gel as quickly after pouring as possible.
· Combs do not need to be deep in the gel, just far enough into the gel to form wells
5) Wet 2 chemwipes and place them on either side of comb. Let polymerize for 1 hour or overnight
To store in fridge overnight, keep wet chemwipes in bag and tape bag closed
WHILE GEL POLYMERIZES, prepare samples
Step Two: Sample preparation
1) Shuck oysters and remove gill tissue.
2) Rinse with dH20 and dry with chemwipes.
3) Dilute gill tissue in KGB buffer
· Dilution: 1 gm oyster gill/ 1000 uL KGB
o Weigh tissue first, then add an appropriate concentration of KGB buffer
· .4179 g tissue/4,179 uL KGB
o Homogenize sample/KGB
· Diluted sample can be aliquoted into centrifuge tubes and stored at –80 degrees.
o I made aliquots of 75 uL
1) Add equal volumes of KGB + sample mixture with 2X Sample Buffer to centrifuge tubes
· Be sure that Sample Buffer is fresh. B-mercaptoethanol degrades over time. . . do smell test.
2) Vortex
3) Boil in a bath of boiling dH2O for 5 minutes
· Be sure that the lid to the centrifuge tube is locked close
· Do not submerge the tube. Put it only halfway in water
4) Remove sample from water bath and wipe off lid

RUN GEL
1) Gel set up (helpful hints)
· Make sure that the gel is square in the apparatus and that the clamps are flush on both sides.
· There must be either two gels running or one gel and one “blank” at all times. The “blank” must be set up identical to the actual gel.
· Tighten the gels into place
o Make sure that they are properly situated by pouring a little running buffer into the electrode area between the two gels. If the buffer leaks out, then the gels are not secured tightly.
2) Remove comb from gel and load 10 uL of sample into the first 8 lanes and the sample ladder into the 9th lane.
· Use extra long pipette tips to load the gel
3) Hook gel box up to a power source and run for 90 minutes at 210 Volts. Record starting and ending amps.

GEL TRANSFER

1) Soak transfer pads in transfer buffer for the last 30 minutes that the gel is running. Add nitrocellulose membrane for the last 10 minutes.
· When cutting the transfer pads and nitrocellulose membrane I use a clean gel plate to estimate the size of the pads/membrane.
· Do not touch the membrane with your fingers. Always use forceps.
2) Stop polyacrylamide gel about 30 minutes after the ladder reaches the end of the gel.
3) Place gel into transfer buffer to equilibrate it
4) Create gel sandwich: pad-gel-nitrocellulose-pad and assemble so that the membrane is closest to the “red” side of the power unit (in our lab, the unit runs from bottom to top).
5) Run at 10 volts for 70 minutes.
6) Record starting and ending amps
7) Place “gel sandwich” back into transfer buffer.
8) Cut the bottom right corner of both the gel and of the membrane for alignment purposes
9) Place the gel in a pipette box and cover with Coomassie Blue in the fume hood and let soak overnight.
10) Place the nitrocellulose in a different pipette box and cover with Ponceau in the fume hood. Let sit for 5 minutes.
11) While it sits, prepare dry milk in TBST (5 g/100 mL)
12) Pour Ponceau into a bottle for reuse.
13) Rinse nitrocellulose membrane with distilled water and photograph and/or photocopy
14) Pour TBST and Milk over the membrane and store in the refrigerator overnight.

PROBE GEL WITH ANTIBODIES

1) Rinse gel for 10 minutes, 3x with TBST
2) Add Primary antibody to TBST
· 1:1250 dilution
· Helpful hints
o We dilute the antibody for storage at –80 to avoid freeze thaw cycles
o Be sure to include the original dilution factor into later dilutions
3) Rotate primary antibody at 57 RPM’s for 2 hours
4) Rinse 3x at 10 minutes each
5) Add secondary antibody to TBST
· 1:3000 dilution
6) Rotate at 57 RPM’s for 1 hour
7) Rinse 3x 10 minutes each
While antibodies rotate/incubate, dry polyacrylamide gel
· soak in destain for 2 hours
· Rinse with water
· Soak in drying solution for 30 minutes
· Use OWL gel drying kit
o Soak cellophane in dH20 for 2-3 minutes.
o Place cellophane on solid acrylic plate
o Put gel on top of cellophane
o Drop some drying solution on top of membrane (~2 mL)
o Place cellophane on top of membrane, taking care to minimize bubbles.
o Let dry on bench overnight.

PHOTOGRAPH MEMBRANE

1) Shake off all excess TBST from membrane
2) Mix chemiluminescent reagents according to instructions
3) Apply to membrane and incubate in the dark for 1 minute
4) Shake off excess chemiluminescent
5) Focus camera
6) Take one picture with light to photograph the ladder
7) Close door and take a picture at an exposure time of 2 minutes.
8) Take another picture at 5 minute exposure.
9) To adjust the image, under Edit menu, select Display range
· Move the bar on far right to the left until bands appear.
· Invert the image to get black bars on white image.
10) Save and print all images.



RNA Isolation

(2/11/09 LC)
*important to use extreme sterile technique when handling RNA

1. Turn on heating block to 55C

2. Add 500uL of TriReagent *caution: contains phenol – use in hood!* to a 1.5mL snap cap tube. Store on ice.

3. Using a clean razor blade, 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.
----- Can stop here and freeze sample at -80

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.

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.


Don’t forget to quant RNA yield before reverse transcription!!

Reverse Transcription

Standard Operating Protocol (SOP)

Written 20150702 by Sam White.
Reagents:
Personal Protective Equipment (PPE):
  • Gloves
Equipment:
  • Pipettes (10 - 1000uL)
  • Filtered pipette tips
  • 0.5mL snap-cap microfuge tubes (Genesee: 22-178A)
  • Sterile 1.7mL snap-cap microfuge tubes (Genesee: 22-281S)
  • Thermal cycler, water bath, or heating block capable of 37C OR 42C.
  • vortexer
  • ice

Procedure

Total Time: ~ 1.5 - 2.0hrs
Cost/sample: ~ $1.50
IMPORTANT: A single reaction volume = 25uL. The volume of RNA, primer(s) and M-MLV RT used in this protocol are variable and will be specific to your current experiment. The directions below apply to a reaction using 1ug of total RNA. You may need to make changes to accommodate your own conditions.
  1. Read the manufacturer's protocol.
  2. Read this protocol.
  3. Verify sufficient quantities of reagents and samples before beginning.
  4. Wear clean gloves.
  5. Thaw all RNA and reagents on ice. Prepare all reactions on ice.
  6. Transfer 1ug of RNA to 0.5mL snap cap tubes or PCR plate. Adjust volumes of individual samples to 17.75uL with H2O.
  7. Add 0.25ug primer per 1ug of RNA in sample (= 0.5uL of Promega oligo dT Cat#C1101 in this example). Total volume (RNA + primers) should equal 18.25uL.
  8. Heat samples at 70C for 5 min in thermal cycler, heating block, or water bath.
  9. Place samples on ice IMMEDIATELY.
  10. Make Master Mix:
    Per Reaction
    • 5 uL 5x Buffer (M-MLV RT Buffer)
    • 1.25 uL 10mM dNTPs
    • 0.5 uL M-MLV RT per ug of RNA
  11. Mix well by flicking; do not vortex.
  12. Add 6.75uL of master mix to each reaction.
  13. Mix by pipetting; do not vortex.
  14. Incubate @ 42C for 1hr for oligo dT primers OR @ 37C for random primers.
  15. Heat inactivate @ 95C for 3 min.
  16. Spot spin and store @-20C.


Extraction of DNA from Sediments for PCR Amplification

(Jennifer L. McLarnan 6/2000, revised from James P. Gray method 5/96)

Extraction

1) Make lysis buffer and heat at 55C for 60 minutes. Lysis buffer must be made just prior to use and heated for 60 minutes before using. Use buffer warm.
Buffer: -500mM NaCl (100ul of 5M per ml)
-50mM EDTA (100ul of 0.5M per ml)
-50mM Tris pH8.0 (50ul of 1M per ml)
-4% SDS (160ul of 25% per ml)
-npH20 (590ul per ml)
2) Into a 2.0ml beadbeater tube weigh out the following (in order):
2.0g 0.10mm zirconia/silica beads
400.0mg sediment
900.0ul lysis buffer (55C)
3) Homogenize samples in beadbeater for 1.0min on the “homogenize” setting.
4) Incubate tubes in 70C waterbath for 60 min.
5) Centrifuge samples at 13,000rpm for 2 minutes using the Eppendorf 5415C centrifuge.
6) Using a wide bore pipet tip, transfer supernatant to a new 1.7ml tube.
Incubate samples 15.0min on ice.
7) Centrifuge samples in the 5415C centrifuge that has been pre-chilled to 4C. Centrifuge at 14,000rpm for 5.0min at 4C.
8) Clean supernatant with an acid washed PVPP column, using one column per sample.
9) Add 2 volumes of –20C 100% ethanol.
10) Invert tubes 20 times.
11) Incubate tubes at –20C for 15minutes.
12) Centrifuge samples at 14,000rpm (4C) for 5.0 minutes.
13) Discard supernatant and wash pellets once with 1.0ml –20C 70% ethanol.
14) Air dry pellets.
15) Resuspend in 50ul TE (pH 7.5).
16) Proceed immediately to Purification.

Purification

1) Quantitate the extracted DNA using the GeneQuant II.
2) Run the samples on a 1% Agarose gel (1x modified TAE) at 5 V/cm.
Add guanosine to the gel and buffer and post-stain the gel with ethidium bromide.
3) Excise the large MW band using a clean razor blade for each sample.
4) Isolate the DNA from the agarose using Bio 101 GeneClean III Kit with Spin. Follow directions for purification from agarose. Bind the DNA to the glassmilk by incubating the tubes at room temperature for 15-20 minutes while providing gentle agitation or occasional flicking. After wash steps, centrifuge empty columns 15 minutes at 14,000rpm to dry pellets. Elute samples once using GeneClean Elution Solution. The elution volume used should be equal to 3 times the glassmilk pellet. To elute, incubate at room temperature for 15-20 minutes (with occasional flicking) prior to centrifuging.
5) Quantitate the DNA with the GeneQuant II or by running aliquots of DNA on a 1% Agarose gel and compare to High Mass DNA Ladder (Gibco BRL).
6) Proceed immediately to PCR Amplification.

PCR Amplification

1) Set up PCR reactions per separate protocol.
2) Run thermocycler program “16S-JLM” on the MJ Research Minicycler.
3) After cycle is complete verify product by running 5ul on a 0.5% Agarose gel (1xTBE).

Setting up a Restriction Enzyme Reaction(New England Biolabs, Edited by R.E. Strenge 6/04)

A "Typical" Restriction Digest
Most researchers follow the general rules that 10 units of restriction enzyme is sufficient to overcome variability in DNA source, quantity and purity. Generally, 1ul of enzyme is added to 1 ug of purified DNA in a final volume of 50 ul of the appropriate 1X NEBuffer followed by incubation for 1 hour at the recommended temperature. If an excess of enzyme is used, the length of incubation can often be decreased to save time. Alternatively, you can productively digest with fewer units of enzyme for up to 16 hours with many restriction enzymes.
Choosing the Right Enzyme
Obviously the DNA to be digested must contain a recognition sequence for the restriction enzymes chosen. Restriction enzymes with shorter recognition sequences cut DNA more frequently than those with longer recognition sequences. Assuming a 50% G-C content, a restriction enzyme with a 4-base recognition sequence will cleave, on average, every 44 (256) bases compared to every 46 (4096) bases for a restriction enzyme with a 6-base recognition sequence. Cleavage by a restriction enzyme produces either cohesive (having either a 5´ or 3´ single-stranded protrusion) or blunt-ended (no single-stranded protrusion) fragments. Cohesive fragments can be subsequently ligated to other restriction fragments if their single-stranded protrusions or "overhangs" are compatible. All blunt-ended fragments can be ligated to each other. See Compatible Cohesive Ends and Recleavable Blunt Ends.
Enzyme
Restriction enzymes should be kept on ice when they are not in the freezer. The enzyme should always be the last component added to a reaction (reaction components should be mixed prior to addition of enzyme). The number of units added to a reaction must be adjusted to the varying cleavage rates of DNA substrates. For example, supercoiled plasmids and agarose-embedded DNAs generally require more than 1 unit/µg to be cleaved completely. More information about cleavage of plasmid DNA and agarose-embedded DNA.
DNA
The preparation of DNA to be cleaved should be free of contaminants such as phenol, chloroform, alcohol, EDTA, detergents, or excessive salts, all of which can interfere with restriction enzyme activity. DNA methylation is also an important element of a restriction digest. More information about Effect of CpG Methylation on Restriction Enzyme Cleavage and Dam and Dcm Methylases of E.coli.

Reaction Buffer
New England Biolabs provides a color-coded 10X NEBuffer with each restriction enzyme to ensure optimal (100%) activity. The buffer should be used at 1X concentration in the reaction. Some restriction enzymes require bovine serum albumin (BSA) at a final concentration of 100 µg/ml for optimal activity. When required, BSA is supplied as a 10 mg/ml (100X) stock and should be added to the reaction mixture. Restriction enzymes that do not require BSA for optimal activity are not adversely affected if BSA is present in the reaction. More information about NEBuffers.

Reaction Volume
By definition, 1 unit of restriction enzyme will completely digest 1 µg of substrate DNA in a 50 µl reaction in 60 minutes. This enzyme : DNA : reaction volume ratio can be used as a guide when designing reactions. Smaller reaction volumes are more susceptible to pipetting errors. To keep glycerol concentration at less than 5% in a reaction, the restriction enzyme, which is supplied in 50% glycerol, should not exceed 10% of the total reaction volume.

Mixing
An extremely important, yet often overlooked, element of a successful restriction digest is mixing. The reaction must be thoroughly mixed to achieve complete digestion. We recommend gently pipetting the reaction mixture up and down or “flicking” the reaction tube. Follow with a quick (“touch”) spin-down in a microcentrifuge. Do not vortex the reaction.

Incubation Temperature
The recommended incubation temperature for most restriction enzymes is 37°C. Restriction enzymes isolated from thermophilic bacteria require higher incubation temperatures ranging from 50°C to 65°C. More information on the activity of thermophiles at 37°C.

Incubation Time
The unit definition of our restriction enzymes is based on a 1 hour incubation. Incubation time may be shortened if additional units of restriction enzyme are added to the reaction. Conversely, longer incubation times are often used to allow a reaction to proceed to completion with fewer units of enzyme. Refer to the specific information about enzyme survival times in a reaction.

Stopping a Reaction
If no further manipulations of the digested DNA are planned, the reaction can be terminated by adding a stop solution. At NEB we use the following stop solution: 50% Glycerol, 50 mM EDTA (pH 8.0), and 0.05% bromophenol blue (10 µl/50 µl reaction). If further manipulations of the digested DNA are required, heat inactivation (raising the temperature to 65 or 80°C for 20 minutes) is the simplest method of stopping a reaction. Since this method does not work for all restriction enzymes, refer to the heat inactivation chart. Phenol/chloroform extraction is another means of inactivating a restriction enzyme.

Storage
We recommend storage at –20°C for most restriction enzymes. For a few enzymes, storage at –70°C is recommended for periods longer than 30 days. Please refer to the enzyme's technical data sheet or catalog entry for storage information. 10X NEBuffers and concentrated BSA should also be stored at –20°C. BSA should not be mixed directly into NEBuffers and then frozen because the BSA may precipitate.

Stability
All enzymes are assayed for activity every 1-2 months; the most recent assay date is given on the label attached to each vial of enzyme. After thirty years of experience with restriction enzymes, we have found that most are very stable when stored at –20°C in the recommended storage buffer. Exposure to temperatures above –20°C should be minimized whenever possible.

Control Reactions
If you are having difficulty cleaving your DNA substrate, we recommend the following control reactions. Incubate experimental DNA without restriction enzyme (degradation of DNA indicates contamination in the DNA preparation or reaction buffer) and control DNA (DNA with multiple known sites for the enzyme, e.g. lambda or adenovirus-2 DNA) with restriction enzyme to more accurately judge whether or not the reaction went to completion. If the control DNA is cleaved and the experimental DNA resists cleavage, the two DNAs can be mixed to determine if an inhibitor is present in the experimental sample. If an inhibitor (often salt, EDTA or phenol) is present, the control DNA will not cut after mixing.

Shipping Samples (in ethanol) for Central Histology Facility

(R. Strenge)
Histology samples from the Friedman lab are placed in cassettes and stored in 70% ethanol until further histology processing. Ethanol is considering a hazardous material. Ground shipping allows for limited quantities to be offered for shipment exempt from labeling and strict packaging requirements. These guidelines must be followed for proper shipping:
  • Wrap cassettes in paper towels and place in a ziplock bag.
  • Add 30 ml of ethanol to the paper towels. It is OK to use multiple containers if there are many samples. (No more than 1 litre total per package).
  • Place ziplockbag into another ziplock bag and place these in one of the small screw cap, plastic containers we purchased specifically for histology samples.
  • Place the lid is on tight!
  • Seal lid with 2 layers of parafilm.
  • Place in a styrofoam shipping cooler. (This helps keep the samples cool and moist.)
  • Add enough absorbent material (Sphagsorb or some from previous shipments) to equal the volume of ethanol being shipped.
  • Add peanuts if necessary to keep container from moving around.
  • Add instructions for CHF to cooler.
Have packaging inspected and approved by Lisa or Sammi.

  • Put lid on cooler and seal with lots of tape.
  • Place cooler in outer cardboard box.
  • Place two “This End Up" labels on opposing sides of the container indicating proper package orientation.
  • Identify shipment with the “Dangerous Goods In Excepted Quantities” sticker. Ethanol is Class 3, UN1107.
  • Ship via UPS ground.

Total RNA Extraction (Robert's Lab Protocol)


Manufacturers' Protocol - MRC
1. Turn centrifuge on to cool to 4C
2. Clean Homogenizer - Rinse in DEPC water in 50 ml falcon tube (3x – 3 separate tubes)
3. Get sample and thaw enough to get out of container
4. Measure weight of sample
5. Take sample out (screw out and use forceps) and chop up with sterile razor blade
6. Put tri-reagent (stays on ice when not using & is light sensitive) into 50 ml falcon tube (or smaller tube depending on size of sample) – for a 0.7 gram sample I used 7 ml of tri-reagent. Note: in 50 ml falcon tube need at least 3 ml of tri-reagent to get it to work
7. Add sample
8. Keep on ice
9. Blot homogenizer with paper towel to remove excess water
10. Homogenize sample (don’t leave off ice for too long)
11. Homogenize until sample is in solution
12. Transfer all or part (I kept 6 ml) of mixture into a 13 ml tube (only add up to 7 ml)
13. Let sit for 5 min at RT
14. Rinse homogenizer in DEPC water (same tube used to clean in the beginning)
15. Add 0.2 ml of chloroform (under hood, open only in hood, pour into glass beaker first) per 1 ml of tri-reagent
16. Cover & shake
17. Let sit for 15 min at room temp
18. Change gloves
19. Spin at 12,000 x g (11,500 rpm) for 15 min at 4C
20. Transfer aqueous (top) phase to fresh tube (top layer has RNA in it – bottom layer has DNA and proteins)
21. Add 3 ml iso. (2 – Propanol, under hood) to precipitate out the RNA
22. Cap and vortex
23. Let sit at RT for 10 min
24. Put waist with tri-reagent etc. in tri-reagent bottle in fume hood
25. Clean up homogenizer and put away (put in 50 ml falcon tube with 5-7 ml 30% H2O2 and up to 40 ml with DEPC water
26. Spin at 12,000 x g (11,500 rpm) for 15 min at 4C
27. Remove supernatant (I want the pellet – RNA)
a. Get glass beaker and paper towels (small stack)
b. Pour off supernatant into beaker and place tube upside-down on paper towels
c. Note: do not rock the tube back and fourth or will loosen pellet
28. Add 1 ml 75% EtOH in DEPC water per 1 ml of tri-reagent added in beginning
29. Cap & move – rock back and fourth to loosen pellet – vortex if necessary
30. Spin 11,500 x rpm for 5 min at 4C
31. Remove supernatant again & put on paper towel – be much more careful to make sure pellet does not slip out
32. Spot Spin – turn on centrifuge, let go up to about 1000 rpm then shut off – note: place pellet facing upward
33. Use filter pipette tips to remove excess EtOH
34. Turn upside down on paper towels
35. Wait 10 mins
36. Depending on size of pellet add dnase free water to the tube – if taking to mRNA always use 500 ul (I used 500 ul for the ovary – large pellet, and 250 ul for the muscle)
37. Dissolved into solution by pipetting
38. Put in 1.5 ml tube
39. Put on ice
40. Spec to determine how much RNA you have

DNA Extraction for Water Samples on Filter paper


Handling after/during sampling:
1. Have or place filter in boilproof microcentrifuge tube, unroll filter so that more of the inner (or upper when filtered) area is exposed.
2. Freeze filter in -80ºC freezer.
Need for DNA extraction:
Microcentrifuge tubes for DNA aliquot
Sterilized (bleached, rinsed WELL, UV’ed) pestle grinder-1 per sample
Sterile low TE
10% bleach bath
DNA Extraction:
3. Add 300 ml low TE and vortex well for ~30 seconds, trying to get the filter paper as washed over as possible.
5. Place in heat block set at 95ºC for 5 min.
6. Vortex in Disrupter Genie for 2 min.
7. Centrifuge at max speed for 15-30 seconds to get buffer off lid.
8. Using a sterile pestle, mash the filter into the bottom of the tube and mash/grind into it a bit. (Put the now contaminated pestle into 10% bleach bath immediately after removing it to avoid contamination.)
9. Place the tube again in the heat block at 95ºC for 5 min.
10. Centrifuge at max speed for 15- 30 seconds.
11. Store at 4 ºC until QPCR analyses and then store the stock tube with filter into -20 ºC long term.

50 ml
100 ml
200 ml
500 ml


50 ml
100 ml
200 ml
500 ml
AUTO-POW
45.8
91.6
183.2
458

AUTO-POW
45.8
91.6
183.2
458
FBS
5.2
10.4
20.8
52

FBS
2.5
5
10
25
L-glutamine
0.5
1
2
5

L-glutamine
0.5
1
2
5
Sodium Bicarb Buffer
0.6
1.2
2.4
6

Tris Buffer
0.7
1.4
2.8
7
Penn-Strep (ml)
0.5
1
2
5

Penn-Strep (ml)
0.5
1
2
5
Gentamycin Sulfate (ml)