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Wayne:Laboratory Protocols
Lab safety
General guidelines:
- You must always wear closed-toes shoes and a lab coat when in the lab!
- Always use gloves when working in the lab but DO NOT WEAR GLOVES OUTSIDE OF THE LAB (e.g. hallway, elevator, etc).
- Pre-PCR is carried out in room 4146. All post-PCR activity takes place in room 4162. Discard gloves when moving between rooms and keep all equipment in their designated rooms. AVOID moving things back into the post-PCR room 4162. These precautions are to avoid contamination problems.
- If you use the last of the supplies (such as tips or gloves), please restock these items.
- When stock solutions, kits, gloves, Kimwipes, etc runs out, it is your responsibility to make sure these get replaced. If more needs to be ordered, please notify Sarah.
- At the end of the day, remove all of your stuff from the bench you were using and return all equipment to their appropriate places, washing glassware with deionized water and hanging on the rack to dry.
- Empty benchtop trash into the appropriate receptacles (i.e. biohazard containers, general laboratory trash contains, recycle bins, etc).
- Make your own aliquot of stock solutions (to avoid contamination).
- Label and date EVERYTHING.
- If you borrow reagents, you must return them.
- Know where the first-aid kit is located.
- Do not prop open the laboratory doors. Many of the researchers keep personal belong- ings in the office/lab. We want to prevent theft and follow Environmental Health and Safety guidelines.
- Please turn off equipment if it is not in use. Especially hot plates! Make sure they are off when you are done using them.
- If you uncomfortable at all with any laboratory technique, please do not attempt the procedure and ask for guidance.
Your supervisors (graduate student or post-doctoral researcher and Dr. Wayne) are responsible for you. Please make every effort to be a good laboratory citizen. The Wayne lab is filled with friendly and knowledgeable people. Please ask questions when you are at all uncertain about your technique, procedure, or project. Please make every effort to attend our weekly laboratory meetings, as this is an important forum for meeting the lab members and keeping up-to-date on issues in the lab (i.e. new techniques, current research projects underway, presenting your own work, etc.). This is also the time for the rest of the Wayne lab to meet you, so they are familiar with you and are able to provide help if needed.
Visit UCLA EH&S for more safety information
Visit Laboratory Safety Fundamental Concepts Online Refresher to complete your annual lab safety online refresher
Visit Safety Classes to sign up for a safety training class if you are new to our lab
Handling hazardous materials
Be extra careful when handling reactions or waste containing hazardous chemicals:
- Polyacrylamide is a neurotoxin and wear gloves. As a liquid it is particularly nasty. Wear nitrile gloves when using it, as acrylamide goes through regular latex and vinyl gloves.
- Ethidium Bromide (EtBr) is carcinogenic and wear gloves. Any waste or contaminated material (including stained gels) must be placed in the labelled containers under the UV camera.
- Always work in the hood when handling organic solvents, like phenol and chloroform, and place waste in appropriately labelled containers.
- Formamide is carcinogenic and wear gloves. Be careful not to breathe near it, as it aspirates easily.
Go to UCLA Online Tag Program to generate a hazardous waste tag
Go to Chemffx for Material Safety Data Sheets (MSDS)
Recipes for the general-use buffers
50X Tris-acetate-EDTA (TAE) (makes 1L)
1. Obtain a bottle with at least 1L capacity
2. Measure out 242 g of solid Tris Base and add to bottle
3. Add 600 ml distilled water
4. Place beaker on stirring block, add stir bar
5. Stir until Tris is dissolved
6. Add 100 ml 0.5M EDTA
7. Add 57.1 ml Glacial acetic acid (wear eye protection and use caution)
8. Add water until total volume is 1L
9. Remove stir bar with magnet
10. Store at room tempurature
5M Sodium Chloride (NaCl) (makes 250mL)
1. Obtain a beaker with at least 250mL capacity
2. Measure out 73.05g of solid sodium chloride (NaCl) and add to beaker
3. Add distilled water up to total volume of slightly less than 250mL
4. Place beaker on stirring block, add stir bar
5. Stir and heat slightly
6. Add water until total volume is 250mL and sodium chloride is totally dissolved
7. Remove stir bar with magnet and place solution in a 500mL bottle with a loosely fitted cap
8. Autoclave (liquid cycle: ~20 mins)
9. Once cooled, run solution through sterile filter system
- Obtain a short length of plastic hose to connect vacuum line to filter unit
- Make sure receiving unit (bottom bottle) and filter unit (top half) are held together tightly
- Slowly turn on vacuum and keep at a low pressure
- Pour solution into filter unit and allow to drain into receiving unit
- Turn off vacuum, remove filter unit, screw on cap
10. Label container (common chemical name, concentration, date, your initials)
10X loading dye(makes 50mL)
1. Obtain a 50ml Falcon tube
2. Add 19.5mL glycerol
3. Add 1mL 0.5M EDTA
4. Add ~39mL distilled water top bring volume to 50mL
5. Close lid and shake
6. Transfer 10mL into 15mL Falcon tube
7. Measure 0.025g of Orange G (>100bp*), Bromophenol blue (400-500bp*), AND/OR Xylene cyanol (10,000-4000 bp*)
- based on 0.5-1.5% agarose gel
8. Add dye(s) to Falcon tube
9. Close lid and shake
10. Store in refrigerator
11. Label container (common chemical name, concentration, dye(s) added, date, your initials)
1M Tris(-Cl), pH 8 (makes 250mL)
1. Obtain a beaker with at least 250mL capacity
2. Measure out 30.275g of Tris base and add to beaker
3. Add 200mL distilled water
4. Place beaker on stirring block, add stir bar
5. Stir and heat slightly to dissolve Tris base
6. Calibrate pH meter with pH 7.0 buffer and pH 10.0 buffer
7. Measure pH of solution
8. Add 9mL of concentrated HCl (be very careful, this acid is highly caustic) and measure pH
9. Continue adding HCl dropwise until pH measures 8.0
10. Add distilled water to total volume of 250mL
11. Remove stir bar with magnet and place solution in a 500mL bottle with a loosely fitted cap
12. Autoclave (liquid cycle: ~20 mins)
13. Once cooled, run solution through sterile filter system
- Obtain a short length of plastic hose to connect vacuum line to filter unit
- Make sure receiving unit (bottom bottle) and filter unit (top half) are held together tightly
- Slowly turn on vacuum and keep at a low pressure
- Pour solution into filter unit and allow to drain into receiving unit
- Turn off vacuum, remove filter unit, screw on cap
14. Label container (common chemical name, concentration, date, your initials)
Qubit quantification of double-stranded DNA
Qubit gives a more accurate reading of the amount of dsDNA in a sample. Because we are using a High Sensitivity (HS) kit, your samples should be 1-100 ng/uL to ensure accuracy. You can use anywhere from 1-10uL of sample in each tube, always with a total volume of 200μL. We use 2μL to save sample and avoid potential pipetting errors of using 1μL.
- The dye used in Qubit is light sensitive. Keep dye tube covered in aluminum foil. Put sample tubes with dye in tube rack covered with aluminum foil or in a draw when possible.
- Qubit dsDNA HS Assay Kit
- HS buffer
- HS standard #2 (10ng/μL or 100ng/μL)
- Quant-it dsDNA reagent (light sensitive dye)
- Qubit Assay tubes
- Tinfoil
1. Thaw High Sensitivity buffer, light-sensitive dye, standard and samples to room temperature (or use room temperature stock).
2. Cover a tube rack in aluminum foil for storing samples (or place in a drawer).
3. Pull Qubit tubes for number of samples plus two for the standards.
4. Label tubes STD-1 and STD-2, and label one tube per sample you will quantify. Put in tube rack covered in aluminum foil (or drawer).
5. In a sterile 1.5 mL tube covered in tinfoil mix the following. You should master mix this for the number of samples plus three to include standards and extra.
- Working solution per sample:
- 200μL of HS Buffer
- 1μL of Reagent Dye
6. Mix thoroughly by pipetting or vortexing. Centrifuge briefly.
7. Make Standards:
- STD-1 (this is the blank)
- Add 200μL working solution
- STD-2 (100ng/μl STD)
- 190ul working solution
- 10ul of 10ng/ul standard
- Mix thoroughly by pipetting or vortex briefly and spin down.
8. Make sample tubes:
- Aliquot 199μL of working solution into each sample tube
- Add 2μL of sample in each tube
- Mix thoroughly by pipetting or vortex briefly and spin down.
9. Make sure all liquid is collected at bottom of tubes, drops on the side are not acceptable.
10. Keep all tubes in covered tube rack and transport to core or 2nd floor in the Alfaro lab.
Qubit Machine
1. Tap screen to turn on.
2. Tap DNAhs.
3. Read new blanks and then follow instructions on screen.
- Place STD-1 tube into machine, close lid, and press measure.
- Place STD-2 tube into machine, close lid, and press measure.
4. Begin measuring samples.
- Place sample in machine, close lid, press measure.
- Record number on front screen. It should be in ng/mL. There are options for changing the units to ng/μL if you prefer.
- For ng/µl, press "Calculate stock concentration" and choose appropriate units, making sure the number of µl is what you added (1 or 2 µl).
5. At the end of process, measure STD-2 (same tube) again. It should be around 500 or within 10ng of 100ng.
- If any of the readings for the standards are not expected, repeat the whole process starting at step 1.
Calculations (if you didn't choose "Calculate stock concentration" above):
(Reading of individual sample X 100)/μL of DNA added= ______ng/μL
Nanodrop 2000 quantification of DNA
Please use your own materials (pipette, pipette tips, Kimwipes, etc). Start NanoDrop 2000 software and select analysis method (e.g. Nucleic Acid for DNA). Carefully apply a droplet of water (1-2µL) to the pedestal to clean and initialize the instrument. Use a Kimwipe to clean the pedestal and the top in between readings. At the prompt, name the results file and save it in the "My Documents" folder. Apply 1-2µL to the pedestal and press the "Blank" button (AE buffer, TE buffer, water, etc). Clean the pedestal and apply 1-2µL of sample, then press the "Measure" button. After measuring, the concentration and other data appear in the software window. Continue measuring remaining samples. After measuring, view the results table.
GenomiPhi DNA amplification protocol
1. Retrieve sample buffer and reaction buffer (thaw at room temp); keep enzyme in -80°
2. Label PCR tubes for each individual
3. Bring template DNA into main lab on ice
4. Add 9µl Sample buffer to labeled PCR tube
5. Add 1µl DNA template
6. Use GARY2 WGA directory 95°C for 3min
- Heat to 95°C for 3 min; Cool to 4°C forever
7. While waiting, make master mix (put enzyme on ice)
8. Add 9µl reaction buffer for each sample into new 0.65ml tube
9. Add 1µl enzyme for each sample to reaction buffer
10. Retrieve samples from thermocycler
11. Add 10 µl master mix (reaction buffer + enzyme) to cooled samples
12. Use Gary WGA directory WGAv2
- Incubate at 30°C for 90 min; Heat to 65 °C for 10 min; Cool to 4°C forever
13. Retrieve samples from thermocycler; spin down
14. Add 80µl of EB buffer to cooled samples (found in Qiagen MinElute PCR Purification Kit)
15. Transfer to 1.7ml labeled epitubes (sample name and WGA)
Polymerase Chain Reaction (PCR): General Introduction
A PCR contains the following necessary reagents:
- PCR-buffer. Salt and pH-stabiliser. User stock of 10x is kept in your box.
- MgCl2. Salt which is required for the Taq polymerase to work. The standard rxn concentration is 1.5 mM (range 1-4). Higher concentrations makes the Taq polymerase less specific and favours amplifications of short fragments. Too much MgCl2 often results in multiple bands. User stock of 25mM is kept in you.
- dNTPs. Free nucleotides (Gs, As, Ts and Cs) of which the artificial DNA copies are made. User stock (10 mM of dNTP Mix, which has 2.5mM of each dNTP) is kept in your box.
- Primers. Single stranded DNA (oligonucleotides), usually the length of 18-30 bp. Primers used for RAPD are normally shorter, 10-15 bp. Stock solutions at 100µM are normally, and user stocks at 10µM are stored in your PCR box.
- Taq DNA polymerase. The enzyme that puts the free nucleotides together. It starts at the 3' end of the primer and uses the complementary DNA strand as a template. User stock of 5 units/µl is kept in your box.
- Template DNA. The source of DNA for the PCR amplification. This could be DNA extracted from blood, skin, feathers, or old PCR products. We use a standard concentration at 25 ng/µl but depending on the organism and protocol, the concentration might need further adjustments (5-100 ng/µl).
In addition to these reagents, you may find that researchers are adding other reagents to their reactions in hope of getting better results. You may consider:
- BSA bovine serum albumin. Prevents binding of DNA to the test tube. Recommended concentration is 10-100µg/ml.
- DMSO dimethylsulfide. Increases linearity of DNA (to prevent formation of secondary DNA structure). Recommended concentration is 1-10% DMSO.
A suggested concentration for a general PCR cocktail:
Reagents | 50μL reaction | 25μL reaction | 10μL reaction |
ddH2O | 30.8μL | 15.4μL | 5.5μL |
PCR 10x buffer | 5μL | 2.5μL | 2μL |
MgCl2 (25mM) | 4μL | 1.5-2μL | 0.8μL |
dNTPs | 4μL | 1.5-2μL | 0.8μL |
Forward primer (10μM) | 2μL | 1μL | 0.4μL |
Reverse primer (10μM) | 2μL | 1μL | 0.4μL |
Taq DNA polymerase | 0.2μL | 0.1μL | 0.08μL |
Template DNA (25ng/μL) | 2μL | 1μL | 1μL |
To set up a PCR:
1. Make sure you have reserved a thermocycler for your usage. The sign-up dry-erase board is in 5202.
2. Create a PCR protocol and calculate how much you need of your master mix/ cocktail (number of rxn + 10%). Don't forget the rxn for the blank (negative control). The master mix should contain everything except the template (but you should add the Taq just before aliquoting).
3. Make sure the PCR bench is clean.
4. Thaw template DNA and all reaction reagents on the PCR bench, except the Taq polymerase which should remain in the freezer until needed (it contains glycerol so it does not need thawing).
5. Program the PCR machine while the reagents are thawing.
6. Mix and spin all reagents and keep on ice (once experienced with the technique, keeping all reagents on ice is not necessary, unless there are unexpected delays).
7. Place an appropriate number of PCR tubes in a tray, and briefly label each row of tubes.
8. Aliquot your desired amount of template DNA prior to adding Taq polymerase to your cocktail.
9. Make the cocktail in an 1.5 ml eppendorff tube (snap cap). Start with adding the ddH20 and save the Taq DNA polymerase to the last step. Mix by using the 200 µl pipette set at 150µl and pipette up and down a few times.
10. Dispense an appropriate volume of master mix into each of the reaction tubes (total rxn minus amount of template DNA).
11. Add the template (change tip between samples!!!!).
12. Fix the lids on the tubes.
13. Bring the tubes over to the thermocyclers in room 4162 (on ice if you haven't programmed them yet--it may take a few minutes for a newly turned on machine to warm up). Start the PCR machine. Select and run your program, and ALWAYS USE A HEATED LID! Load the tubes, close and tighten the lid, then you are ready to go!
Programming the thermocycler (choosing a temperature profile):
The standard PCR starts with a warming up phase of 3 minutes at 94˚C, to make the template DNA single stranded (denatured). Then follows the cyclic phase that characteristically consists of three different steps.
1. 94˚C. This is again the denaturing step that initiates all cycles and is normally set between 30-60 sec.
2. 37-70˚C. The annealing temperature when the primer is allowed to settle on the template DNA. This step is usually set between 30-120sec. The chosen temperature depends on the melting temperature, Tm, of the primer (length and GC-content).
3. 72˚C. The elongation temperature is the optimal working temperature of the Taq DNA polymerase. This step is set between 5-500 sec depending on the length of the desired fragment. A rule of thumb is that the Taq polymerase builds about 1,000 nucleotides per minute.
4. The number of cycles used varies normally between 20-40 depending on the template DNA concentration, quality, length of product, and above all, empirical experience with the focal reaction.
5. The reaction is normally ended by a 10 minute phase at 72˚C. This will allow the Taq polymerase to add a protruding A at the 3' end of the fragments. This step is very important when cloning the PCR fragments by means of TA-cloning.
Recipe for 2% agarose gels (small gel chamber):
1. If a 2% solution is already made up, proceed to step 3 to melt the gel. To make 100 ml of new solution, take weighing paper from the drawer, place it on the balance, tare, and add 2.0 g agarose (on shelf above microwave). Add to flask.
2. Add 100 ml of 1x TAE buffer (in carboy).
- 200 ml 1x TAE with 4g agarose for large gel
3. Place the flsk in the microwave and boil for repeated 30 sec intervals until the agarose has melted (about 2 min). Make sure it doesn't boil over! Use the orange heat-protection gloves when handling the warm flask.
4. Tape the ends of the gel casts and insert combs. Use two combs if you don't need to run the fragments out the full length of the gel--it saves time and materials.
5. When the temperature of the agarose solution reaches 60˚C (you should be able to hold the glass for 5 seconds without burning your hand, but make sure the gel hasn't started to polymerize yet), add 3 µl of SYBR Safe to the agarose solution.
6. Pour the solution into the cast so that the gel is about 2 mm thick. Cover with box and allow it to polymerize (about 15-30 minutes).When polymerized, remove the tape.
Running 2% agarose:
1. Move the gel (still in gel cast) into the gel chamber with wells near the black electrode and remove combs.
2. If necessary, top off the 1x TAE buffer to the fill line of the chamber. The buffer can be reused several times, but should be replaced every second week.
3. Take parafilm and pipet 1-2µl of loading dye corresponding to the number of your samples plus one for a DNA ladder (if needed) (may be useful to use Post-PCR multi-channel pipet).
4. Add 5-20 µl * of the final PCR product to each well (change tips between samples) (may be useful to use Post-PCR multi-channel pipet).
5. Load the first well in the gel with 5 µl of a DNA ladder, and then the samples (PCR rxn and dye). You can reuse tips here--just place the tip in the buffer in the chamber and pipette up and down to flush the residue.
6. Put the lid on the electrophoresis chamber.
7. Turn on the power supply, adjust the voltage (80-100V), and let the gel run for 50-60 minutes.
8. When finished, place the gel on the glass plate in the UV camera box. For our camera box, the UV light turns off when the door is open. However, be cautious about exposing your skin to UV rays for too long: severe burns may develop without protective clothing or eyewear.
9. When finished, carefully dispose of the gel in the regular waste bin. Wipe down the glass with kimwipe and dispose of them in the regular waste bin.
- If you need to clean your PCR product by gel extraction, load your entire PCR product.
- If you need to clean your PCR product by exo-sap or sending to Beckman, load less of your PCR product (5ul) or use a surface tension gel to load even less.
Recipe for 2% agarose gel (Surface Tension):
1. To make 30 ml of new solution, take weighing paper from the drawer, place it on the balance, tare, and add 0.6 g agarose (on shelf above microwave). Add to flask.
2. Add 30 ml of 1x TAE buffer (in carboy).
- 50 ml 1x TAE buffer with 1.0g agarose for large glass plate
3. Place the flask in the microwave and boil for repeated 30 sec intervals until the agarose has melted (make sure it doesn't boil over!). Use the orange heat-protection gloves when handling the warm flask.
4. When the temperature of the agarose solution reaches 60˚C (you should be able to hold the flask for 5 seconds without burning your hand, but make sure the gel hasn't started to polymerize yet), add 1.5 µl of SYBR Safe to the agarose solution.
5. Put binder clips on each side of a comb.
6. Pour the solution onto a clean small glass plate and place comb in after you pour the solution.
7. Cover with box and allow it to polymerize (about 10 minutes).When polymerized, remove the comb.
8. Load 4 µl ladder with 1.5 µl (if needed) and 3 µl PCR product with 1.5 µl dye.
9. Run gel at 100V for 30-35 min.
A general PCR set-up template for keeping notes on the experiment, samples, thermocycler, and master mix.
PCR programs
For 10μL reaction volumes using M13-hybrid primers: [M13MXSTD] on Wayne lab thermocyclers
- 95 ºC for 15 min; (For 25 cycles: 94ºC 30s, 59ºC 90s, 72ºC 60s); (For 15 cycles: 94ºC 30s, 53ºC 90s, 72ºC 60s); 60ºC 30 min; 4ºC forever hold
For 10μL reaction volumes using regular dye-labeled primers (Non-M13): [JPSDMPLX] (a step-down program with hotstart and final annealing temperatures and times recommended by Qiagen)
- 95 ºC for 15 min; (For 12 cycles: 94ºC 30s, 60ºC 90s dropping 0.5°C every cycle, 72ºC 60s); (For 33 cycles: 89ºC 30s, 55ºC 90s, 72ºC 60s); 60ºC 30 min; 4ºC forever hold
A Brief Intro to Primer Design
During PCR, primers are annealed to complementary regions of single stranded molecules (a result of the denaturing step in the PCR cycle program). The primer sequences are then extended by the DNA polymerase in the PCR cocktail. Both of these steps are temperature sensitive and testing (or knowing) the right temperatures for your primers are testable. Primer annealing is often around 50-60°C but can be identified through testing.
Good primer design is essential for successful reactions. Here are some important design considerations:
- Primer Length: Optimal length of PCR primers is 18-22 bp is a general guideline. This length is long enough for adequate specificity and short enough for primers to bind easily to the template at the annealing temperature.
- Primer Melting Temperature (Tm): The temperature at which one half of the DNA duplex will dissociate to become single stranded. Primers with Tm in the range of 52-58°C are ideal, with Tm's above 65°C likely to have secondary annealing. The GC content of the sequence gives a fair indication of the primer Tm.
- Primer Annealing Temperature (Ta): Too high Ta will produce insufficient primer-template hybridization resulting in low PCR product yield, whereas too low Ta may possibly lead to non-specific products caused by a high number of base pair mismatches.
- GC Content: The GC content of primers should be 40-60%.
- Repeats and long runs of a single nucleotide in the primer sequence: Avoid them! But if you need, no more than 4 of any is suggested.
Design your primer using the program Primer3.
Primer testing 'in-silico' using the UCSC Genome Browser (select PCR from the home page) or use this UCSC PCR link!
Microsatellite Qiagen multiplexing PCR setup
These are M13-hybrid primers, with the M13 tag on either the Forward or Reverse primer. Make note for preparing the primer mix below.
Primer Mix (SINGLE LOCUS)
Reagents | 1 reaction (50μL total volume) | 5 reactions (250μL total volume) |
100μM non-M13 primer 1 (stock concentration) | 1μL | 5μL |
2.5μM M13 primer 1 (remember to dilute the 100μM stock) | 2μL | 10μL |
2.5μM M13 dye-label (we mostly use 6FAM but others are available) | 2μL | 10μL |
diH2O | 45μL | 225μL |
Primer Mix (TWO LOCI; total volume=160μL)
Reagents | Locus 1 | Locus 2 | Add: |
100μM non-M13 primer | 8μL | 8μL | |
2.5μM M13 primer | 16μL | 16μL | |
2.5μM M13 6FAM dye-label | 32μL | ||
diH2O | 80μL |
Primer Mix (THREE LOCI; total volume=120μL)
Reagents | Locus 1 | Locus 2 | Locus 3 | Add: |
100μM non-M13 primer | 4μL | 4μL | 4μL | |
2.5μM M13 primer | 8μL | 8μL | 8μL | |
2.5μM M13 6FAM dye-label | 24μL | |||
diH2O | 60μL |
Primer Mix (FOUR LOCI; total volume=160μL)
Reagents | Locus 1 | Locus 2 | Locus 3 | Locus 4 | Add: |
100μM non-M13 primer | 4μL | 4μL | 4μL | 4μL | |
2.5μM M13 primer | 8μL | 8μL | 8μL | 8μL | |
2.5μM M13 6FAM dye-label | 32μL | ||||
diH2O | 80μL |
PCR Mastermix cocktail(1 reaction)
Reagents | 1x |
Primer mix | 1μL |
BSA (10mg/mL; dilute stock using 1μL 100mg/mL stock to 9μL dH2O) | 0.4μL |
diH2O | 2.1μL |
Qiagen Master Mix (pre-made from the kit) | 5μL |
- Add 8.5μL of master mix to 1.5μL DNA
- If you would like to use the Q-solution that comes in the Q-MM kit, reduce the water by 1μL and add in 1μL of the Q-solution.
PCR Mastermix (96 wells + smidegen extra= 105 reactions)
Reagents | 105x |
Primer mix | 105μL |
BSA (10mg/mL; dilute stock using 1μL 100mg/mL stock to 9μL dH2O) | 42μL |
diH2O | 220.5μL |
Qiagen Master Mix (pre-made from the kit) | 525μL |
Add 8.5μL of master mix to 1.5μL DNA
What are M13 primers, anyways?
A process where we can save money by not having to buy primers that are already dye-labeled, we instead add a 16mer sequence tag (we use the M13F –20 sequence 5'-GTA AAA CGA CGG CCA G-3') onto the 5' end of one of the primers (called a M13-hybrid primer). After the first couple rounds of PCR cycles, the 16mer tag gets added onto your copied DNA product. We put a small enough amount of the hybrid primer in the PCR cocktail so that it all gets used up by 20-25 cycles. We also put a small amount of M13F-20 primer (16bp in length) in the mix, and this primer has been dye labeled. We then drop the annealing temperature by 5°C and this allows the much shorter dye labeled M13F-20 primer (16bp) to anneal and be added onto the copied DNA strands. We run another 20 cycles and you end up with dye labeled PCR product that is 16bp longer than than your original primer sized product (good to remember if you are comparing it to results using the regular primer).
This is much cheaper because we can buy the M13-20 dye labeled primer in bulk, we need to use so little, and we do not have to buy unique dye labeled primers for every primer set we wish to use (the M13-hybrid primers cost about $10-$12, instead of $70 - $120 for a dye labeled primer).
For further details on the hybrid primer process see:
Boutin-Ganache I, Raposo M, Raymond M, Deschepper CF (2001) M13-tailed primers improve the reliability and usability of microsatellite analyses performed with two different allele sizing methods. Biotechniques 31 (1), 24-26.
Sending microsatellite products to the Core facility
1. Thaw 1ml of ABI HiDi (Formamide).
2. Prepare mix of HiDi and size standard (Liz 500).
- 9.7μL HiDi and 0.3μL Liz PER SAMPLE.
- 1ml HiDi and 15μL Liz PER 96 SAMPLES.
3. 9.5μL of HiDi/Liz per well in a 96-well plate.
4. Make a 1/20 dilution of PCR product.
- 2μL PCR product to 38ul water.
- Use a dilution plate.
5. Add 2μL of diluted PCR to the 9.5ul HiDi/Liz in each well.
6. Use a sticky lid and centrifuge.
7. Denature at 95°C for 5min (disable heated lid); DENATURE program.
- Place on ice immediately for 5min.
8. Spin down plate and place back on ice.
9. Label plate with name: Wayne_r_<PlateName>, your initials, date.
10. Carry on ice to the Core facility (5th floor Gonda, 5309).
11. Place plate in the bottom shelf of the genotyping refrigerator labeled "Ready to Run with ABI 3700".
12. Plate results will be on the WebSeq webpage, to be genotyped using GeneMapper.
13. Be sure to mark the number of plates you brought to the Core on the Wayne lab form on the freezer.
Microsatellite genotyping using GeneMapper v3.0/3.7
1. Add (Import) samples to project.
2. Set up marker panel and create a Bin Set in Panel Manager; set up bins if alleles are known. This is done in a hierarchical method in the Panel Manager:
- In the left navigation window, click Panel Manager and then "New Kit" (upper far-left button). Name it and select the type of data you want.
- Select on the new Kit you created, then click "New Panel". Name it and press Enter.
- Click on the panel you created, then click "New Marker". Name it, provide marker range, color, and number of repeats, and comments if you choose.
- Add in as many markers you have. I create different Kits for each multiplex combination set. Make sure to name everything consistently so you can link up to them later and you know exactly what it means. Example: name Analysis Method, Panel, Kit, etc all matching the microsatellite multiplex primer mix so you can easily cross-reference them when setting up in GeneMapper.
- Next, select your Kit and in the Bins menu, select New Bin Set. Name it and then you can select it from the drop-down menu. This Bin Set is used in the Analysis Method to again, be clear about how you name your Kits and Panels.
3. Set up Analysis Method specific to your samples in GeneMapper Manager.
- In the GeneMapper Manager, click the Analysis Method Tab, and then click New.
- Follow prompts, then name your Method on "General" tab.
- "Allele" tab: this is where you link your Analysis Method to the Bin Set you just created. You can also click "Use the marker-specific stutter ratio" if your marker stutters.
- If marker is known to stutter, you can the change the stutter ratio (value is a percentage of the time you observe stutter in dataset) and this will avoid "over labeling" of stutter peaks.
- "Peak Detector" tab: click "User Specific (rfu)" and here you can set a Minimum Peak Height requirement; useful to avoid over labeling low-intensity peaks. The higher the value, the higher the intensity requirement for peak calling.
- The remaining tabs have tons of values you can change to better tweak your analysis. I haven't figured them all out yet, but don't let that stop you from having fun!
- Click OK, then Done.
4. Select your Analysis Method, Panel, and Size Standard information on Samples Tab of project, and hold "Ctrl-D" to fill down for all samples.
5. Click "Analyze" (green triangle) button to analyze; Genotype Tab will then be activated.
6. On the Genotype Tabs, each individual is represented by a separate entry for each marker it was run with (if you multiplexed markers). Each run is tested for quality control in many categories. The GQ (Genotype Quality) column is most important and is rated one of three quality levels:
- Green square: good to go!
- Yellow triangle: usually ok to go!
- Red hexagon: bad; it's a no-go!
7. To view all electropheregrams, select all runs in each marker (left navigation window in Genotypes Tab) and click the "Display Plots" button.
8. There are two important buttons in the upper left corner (Peak Selection Mode and Binning Mode buttons). You can view runs and add allele bins here if necessary.
- Peak Selection Mode: Here you can click on any peak, add it as an allele you created previous to the analysis run, delete the allele call, or custom name the allele peak as a "new" allele for that individual peak.
- Binning Mode: Here you can create/delete/edit bins and increase/decrease the marker range as needed. Any changes made are saved when you exit the screen but you will need to RE-analyze for it to be applied to all the other samples analyzed with that marker. Go back to the Samples Tab and click the "Analyze" button.
9. I recommend viewing all runs to double check the calls made by GeneMapper. If you think a run is good, but the program gave it a GQ red hexagon (failed), you can override this by two ways:
- When you make any changes to a run, it automatically replaces quality values with gray triangles. This tells you that you made manual changes to the run.
- You can "right" click on the final GQ value and a prompt appears asking "Override the Genotype Quality of this marker?" Yes will also create the gray triangles.
10. Once you have viewed/edited all the runs for a single marker, you can always sort the entries under the Edit menu. You can also edit the Table Settings in Table Settings Editor to hide some of the excessive columns that normally appear on both the Samples and Genotype Tabs. Make sure to select the Table Settings you created in the drop- down menu in order for it to be applied to your project.
11. To Export your Allele Table, click "Export Table" button and save it in your folder. It is saved as a ".txt" file and easily copy/pastes into Excel. No problem!
12. Congratulations! (as a last note, I recommend re-analyzing all samples for a marker when you add more samples for that marker...keeping allele calls consistent and standardized across PCRs).
- Check that you are viewing the table in the correct table edit selection. Microsatellite data should be under the Microsatellite Default viewing screen, AFLP with the AFLP screen, and so on.
- Check that you have selected the appropriate bin set when viewing alleles. This makes a world of difference.
Roche LightCycler 480 Applications
The LightCycler is a very useful instrument that can:
- quickly genotype small variants (e.g. SNPs, 3bp deletions, etc.) using a high resolution melting (HRM) curve analysis and
- quantify copy number (e.g. of a gene or fragment or enriched genomic library) by quantitative PCR (qPCR)
1. HRM protocol
Primer design is different than of that for microsatellites. The goal is to make as small of amplicon as possible (suggested to be <120bp), as this is best for use in an HRM PCR.
HRM primer design
- Locate SNP (or other genetic variant) and it's position using the UCSC Genome Browser or Ensembl in the most recent genome build
- Obtain about 100 bases of flanking sequence in both directions with the SNP in the middle of the sequence (use the DNA tab to get genomic sequence)
- Use Primer3 online to submit sequence using default primer picking options
- Use the brackets [A] to target the SNP location
- Design as small of amplicons as possible (target product size of 45-80 bases max)
- The SHORTER THE AMPLICON THE BETTER
- Keep decreasing product size until Primer3 can't design primers
- Name the primer set: HRM-chr#.position (example: HRM-chr15.32383555)
Setting up the HRM plate and master mix
Reagents for the HRM Mastermix cocktail (1x) | 1x |
Primer mix (2μM or 4μM - this is up for discussion) | 0.5μL |
Roche MgCl2 | 1.75μL |
diH2O | 2.25μL |
Roche Master Mix (pre-made from the kit and light sensitive!) | 5μL |
DNA 1ng+ | 1μL |
Total | 10.5μL |
1. Thaw reagents on ice and covered, found in the pre-PCR room chest freezer
2. Make 2μM primer mix (1μL of the forward primer + 1μL of the reverse primer + 98μL of diH2O)
3. Make master mix, vortex and spin down
4. Add 9.5μL of the master mix to each well in a 384 well HRM plate (different than PCR plate)
5. Add 1μL of DNA to each well and pipet up and down to mix
6. Seal the plate with a HRM sticky lid and spin down using the centrifuge
7. Carry on ice and covered to the Core facility (Gonda 5309)
8. Ask someone at the Core to run your HRM plate (Know the profile you want to run. They have the std profile saved)
9. When the samples have been sequenced, they will be posted on WebSeq and can be downloaded.
A general HRM set-up template for keeping notes on the experiment, samples, and master mix.
A general method for analysis of the .ixo files you will retrieve from the WebSeq website.
Suggested analysis method:
The HRM analysis is quick and painless. On the desktop, you will find a shortcut icon to the program LightCycler480. After logging in with the correct name and password, look at the top of the screen to find a drop-down menu. Select "Navigator" and then use the "import" button to select the right .ixo file to import. Following this, look in the left navigation window for the button "Analysis". Of the many options that pop up, select "Tm calling" and hit the "check" button. You then have an option to select the whole plate and press the "calculate" button. Then the melting temperatures (Tm) will be provided. You can select the cells you want and in the bottom left part of the screen, individual Tm's will appear. You can select more information regarding each Tm (e.g. width of peak) but this is essentially your genotype! You need to think about matching up genotypes (presence/absence of specific indels or SNPs) with the denaturing temperature. Good luck and happy genotyping!
2. Quantitative PCR
qPCR assays are used to estimate relative enrichment by abundance of target fragment. This is a relatively inexpensive way to determine if the enrichment (or library prep) was successful prior to sequencing.
Remember that if you are specifically investigating copy number variation in the genome, you need a control gene that is known to be of a single copy.
Here is an example PCR cycle for a generic qPCR:
Program Name | Cycles | Analysis Mode | Target (°C) | Hold (hh:mm:ss) | Ramp Rate (°C/s) | Acquisitions (per °C) | Acquisition Mode |
Pre-incubation | 1 | None | 95 | 00:10:00 | 4.8 | --- | None |
Amplification | 40 | Quantification | 95 | 00:00:10 | 4.8 | --- | None |
60 | 00:01:00 | 2.5 | --- | Single | |||
Melting Curve | 1 | Melting Curves | 95 | 00:00:10 | 4.8 | --- | None |
65 | 00:01:00 | 2.5 | --- | None | |||
95 | --- | --- | 5 | Continuous | |||
Cooling | 1 | None | 40 | 00:00:10 | 2 | --- | None |
The PCR cycle program should be applicable to HRM genotyping but as always, the annealing temperature may be altered to match your primer. Another difference is when the acquisition occurs; this is the point in the program at which data (time and temp) is collected. For HRM, this only matters during the final melting curve, whereas in qPCR, this is done repeatedly at the end of every amplification cycle. Check that you are using the appropriate program for your analysis!
In the lab, we use the KAPA kit pretty much as stated in the PDF for quantifying library preps with qPCR.
- Sign up for the qPCR run on the LightCycler!
- Run all samples in duplicate but the 6 standards in triplicate.
Suggested analysis method:
On the desktop, you will find a shortcut icon to the program LightCycler480. After logging in with the correct name and password, look at the top of the screen to find a drop-down menu. Select "Navigator" and then use the "import" button to select the right .ixo file to import. Next, you will need to import (or set up) your plate template, as to tell the program where your standards are of known concentrations in order to use as a ruler for your own samples. To import your template for qPCR, (for this example we will use Rena's template for quantifying HTS libraries using the KAPA kit), go to the Sample Editor and import the file named kappa-plate-setup-2peat-384-14SAMPLES). Open it up and just scan the headers to check that it looks correct, then select "apply". Following this, look in the left navigation window for the button "Analysis". Of the many options that pop up, select "Absolute quantification/2nd derivative max" in order to do a quantification based on your standards, and hit the "check" button. Make sure to change the efficiency in the menu at the bottom to select the "std curve (in run)" option. Also check that "high sensitivity" is selected in the "high sensitivity" menu. Then click the "Calculate" button. You need to then look at the duplicated you created, first checking the standards and that they are consistent. Make sure to check EVERY standard! Then go ahead and begin surveying your own samples. After all is said and done, you can right-click on the Replicate Statistics Table at the bottom to export.
Also, from here, you can use the concentrations and standards to calculate molarity of your libraries and pool based on that (which is a requirement for all HTS sequencing). Here is an example spreadsheet...but remember, MAKE SURE YOU DISCUSS THIS WITH SOMEONE WHO HAS ALREADY DONE THIS PRIOR TO SUBMISSION TO ANY CORE FACILITY. This is an easy place to go wrong, so double check with your colleagues. Most people will dilute libraries down to 10nM before pooling, but this is flexible, depending on your libraries' concentrations. Best of luck!
3. You must sign up to use the Roche LightCycler 480 at Genoseq LightCycler 24 hours in advance!
Dilution calculator
You are welcome to try your own hand at this or find a nice website (e.g. Dilution calculator) that may be helpful for your dilution needs!
A helpful dilution calculator:
C1 x V1 = C2 x V2
- C1: starting concentration with units
- V1: starting volume
- C2: final or desired concentration with units
- V2: final or desired volume
For example:
You have a 2 mg/ml stock solution of BSA, and you use 0.2 ml of that BSA in a total volume of 1 ml.
- You know the stock concentration: 2 mg/mL = C1
- You know the volume of stock solution that you use: 0.2 mL = V1
- You know the final volume of the sample: 1 mL = V2
- You do not know the final concentration of the sample: C2 = unknown (0.2 mL)*(2 mg/mL) = (1 mL)*C2
- C2 = (0.2 mL)*(2 mg/mL) / 1 mL
- C2 = 0.4 mg/mL
The final concentration of BSA will be 0.4 mg/mL.
Targeted sequencing
If you have enough PCR samples for a 96-well plate, please proceed to the Beckman Coulter Sequencing instructions.
PCR products may be cleaned by gel purification (we use zymoclean kits) or by EXOSAP (as follows).
In order to sequence PCR products, excess primers, single stranded DNA, and extra nucleotides (dNTP's) need to be removed. Exonuclease I (Exo) digests single stranded DNA and Shrimp Alkaline Phosphatase (SAP) dephosphorylates free nucleotides making them unavailable for polymerization. Store these reagents at -20 C.
1. Put ice in an ice bucket and retrieve EXO, SAP, and Dilution Buffer tubes from "EXO SAP" box in Post-PCR freezer. Keep EXO and SAP on ice to prevent enzymes from denaturing.
2. Flick or pipette mix EXO and SAP – do not vortex. Make up a master mix containing H2O, Dil. Buffer, EXO and SAP following table below:
EXOSAP Mastermix cocktail (1 reaction in 1 direction)
Reagents | 1x |
diH2O | 1μL |
Dilution Buffer | 0.25μL |
EXO | 0.25μL |
SAP | 0.5μL |
PCR Product | 3.0μL |
TOTAL | 5.0μL |
3. Aliquot 2uL master mix into PCR SnapStrip tubes or 96-well PCR plate.
4. Combine with 3uL PCR product per tube/well. Pipette mix contents.
5. Spin down tubes/plate and run EXO SAP program on PCR machine (~1hr).
6. Thermocycler profile:
a. 37 degrees for 30 min (to perform digestion)
b. 80 degrees for 10 min (inactivate the enzymes)
c. 4 degrees forever
7. Once the EXO-SAP reaction is finished, store your cleaned PCR in the freezer (-20) until ready to set up the BIG DYE reaction (cycle-sequencing).
1. Obtain tube of Big Dye from freezer box in Post-PCR freezer. Keep on ice and cover while thawing (light sensitive).
2. Obtain tube of Seq Saver from refrigerator.
3. Make up master following table below:
BIG DYE Mastermix cocktail (1 reaction in 1 direction)
Reagents | 1x |
diH2O | 3μL |
Seq Saver | 0.5μL |
Big Dye (Light Sensitive!) | 0.5μL |
Primer (3ul) | 1.0μL |
Clean PCR Product | 5.0μL |
TOTAL | 10.0μL |
4. The volume of PCR product can vary from 1.5-5ul depending on how concentrated your PCR product is. Remember to adjust the volume of water if you adjust the volume of PCR product.
5. Aliquot 5uL master mix into PCR SnapStrip tubes or 96-well PCR plate.
6. Combine with 5uL PCR product per tube/well. Pipette mix contents.
7. Spin down tubes/plate and run BIGDYE05 program on PCR machine.
8. Thermocycler profile:
a. 96 degrees for 10 sec
b. 96 degrees for 10 sec
c. 50 degrees for 20 sec
d. 60 degrees for 4 min
- 35x of b, c, d
e. 4 degrees forever
9. Once the BIG DYE reaction is finished, store your cycle sequencing product in the freezer (-20) until ready to send to the CORE.
Sequencing sign up
1. Sign up to sequence at Genoseq CORE.
2. Click on R2R Signup (list) (found on the left hand side) and fill out the required information.
3. Answer YES for request cleaning service!
4. Please bring your reference number with you, remember to check in your samples when you drop them of, and bring back Wayne lab sample racks.
5. Alternatively, you can bring your samples over to the Core and sign up there.
- Carry samples on ice to the Core facility (5th floor Gonda, 5309).
Cloning, colony picking, and PCR
Cloning reaction
1. Thaw cloning reaction reagents. Keep TOPO vector on ice.
Reagents | Full reaction | 1/2 reaction | 1/4 reaction |
Fresh PCR Product | 1μL | 0.5μL | 0.25μL |
Salt Solution | 1μL | 0.5μL | 0.25μL |
Water (add to make) | 3μL | 1.5μL | 0.75μL |
TOPO Vector (keep on ice, add last) | 1μL | 0.5μL | 0.25μL |
Final Volume | 6μL | 3μL | 1.5μL |
2. Mix cloning reaction gently and incubate at room temperature for 5 min (10-15 min for large PCR products (>1kb)).
3. Place on ice.
Transforming competent cells
4. Make sure water bath is at 42°C, shaker is at 37°C, SOC medium is at room temp., plates are at 37°C, prepare 2.0 ml tubes (keep on ice) if cells are to be split (for half or quarter reactions).
5. Thaw tube of competent cells on ice.
6. If using 1/2 or 1/4 reactions, split thawed competent cells amoung chilled 2 ml tubes proportionally (use pipette tip with cut off to minimize shearing of cells). The full reaction volume is 50μL of cells.
7. Add 2μL of cloning reaction to 1 full tube of competent cells, of 1μL to a 1/2 tube of competent cells, or 0.5μL to a 1/4 tube of competent cells.
8. Incubate on ice for 15 minutes.
9. Heat shock the cells for 30 seconds at 42°C in water bath, then place tubes back in ice.
10. Add 250μL of room temperature SOC for full reaction (125μL for half reaction).
11. Make sure tube is sealed well and incubate at 37°C for 1 hour in 200 rpm shaker.
12. While mix is incubating, spread 80μL of 20 mb/ml X-gal solution onto each plate, let incubate at 37°C until ready for plating.
13. Spread 10μL to 50μL of transformation mix into prewarmed plate (we usually plate 2 or 3 amounts, e.g. 20μL and 40μL).
14. Put plate in 37°C static incubator right side up for 20-30 minutes with lid slightly gapped to allow transformation solution to dry, then flip over to incubate until colonies are desired size (typically 24-48 hours).
15. Put in cold room (wrap in parafilm) until colonies are to be picked or grown larger by placing back in incubator.
Proceed to Pick & Boil Step or Growing Selected Colonies and DNA Extraction
Pick & Boil Step
Goal: Use a pipette tip to pick a colony, boil it to lyse plasmid from bacteria, and then PCR it with M13F and R primers to cleanly amplify just the target insert without needed gel band cutout.
Recommend using a new box of sterile pipette 20 or 200 µl size tips (Caution! Don't use tips from a box that has been previously opened and used for other work. If you don't use all the tips, set them aside and label for colony picking use only).
1. Use a 96-well PCR plate and add 30µl of 0.1X TE buffer into each well you plan to use for a picked colony. We recommend using the bottle of molecular grade 50X TE buffer from which we make the 0.1X solution.
2. Using a sterile pipette tip, gently touch the selected colony to get some cells on it, then place the tip into the well of the PCR plate. Gently swirl the tip in the well for a few seconds. After you have picked all the desired colonies and "innoculated" the wells of the plate (a tip should be in a well for at least a minute), then gently remove each tip without dripping any liquid into other wells.
3. Put a flexy silicone lid on the plate and then boil the colonies to spring (lyse) the plasmids from the bacteria. The cycle used should be 95°C for 10 minutes, followed by 4°C forever until you are ready to take the plate out of the PCR machine. Make sure to use the heated lid option!
PCR Reaction Mix & Cycle
You want to put enough plasmid into the PCR reaction to get decent amplification of the insert, but not too much that you have to ultimately cut the insert band out because the plasmid concentration is too high. We have found that 1µl of the boiled colony solution works well to get this result.
Reagents for a 10μL reaction | x1 |
Qiagen Master Mix | 5μL |
BSA (10mg/mL; dilute stock using 1μL 100mg/mL stock to 9μL dH2O) | 0.4μL |
M13 Foward and Reverse primer mix (2μM) | 1μL |
diH2O | 2.6μL |
- add 9μL of PCR cocktail to 1μL boiled colony per reaction
PCR program
- 95 ºC for 15 min; 94°C for 30s; (For 45 cycles: 89ºC 30s, 50ºC 60s, 72ºC 60s); 60ºC 30 min; 4ºC forever hold
Do a checking gel to see if you have an insert amplifying in each sample (if there is a ~200bp band, that means there is no insert; if no band, means there was no or very few cells in the boiled solution). Use 3µl of PCR product and 2µl of loading dye for the checking gel. If clean single band of the expected insert size, do the standard next steps for sequencing (Agencourt/Beckman prep, or if UCLA core – Exosap and Big Dye rxn).
Growing Selected Colonies and DNA Extraction
1. Pick white colonies with sterile pipette tips (label backside of plate to keep track of colonies picked). Blue colonies do not have a PCR product.
2. Put 1.5 ml of LB or TB broth (make sure antibiotic is added into 10 ml tube, pick selected colony with sterile pipette tip and drop in tube. Put cap loosely on tube and put on shaker at 37°C overnight to grow bacteria. Put in cold room after bacteria have grown to desired concentration.
3. Extract plasmid DNA from bacteria using Qiagen MiniPrep kit.
4. Sequence using original PCR primers, or more desirably -M13 (-20) primers, which start ~50-70bp in plasmid (will sequence the beginning of PCR product). Plasmid extraction DNA concentration is high so don't need to add much to sequencing reaction.
Beckman Coulter Sequencing
1. Make sure to use Neptune 96-well plates and tinfoil lids.
- Need to provide:
- 20μL PCR product per well in 96-well plate
- 25μL of primer per well in 96-well plate
2. Log on to Beckman Coulter Sequencing online:
- http://psf.beckmangenomics.com/
- See password sheet in computer room for login and password information
3. Click link for Create a New Project
4. Clink link for Single Pass PCR Sequencing Project under the heading of High Throughput Sequencing
5. Enter a project name
6. Enter the number of samples into 96-Well Qty (# of PCR product plates)
7. Enter Amplicon Size
8. Check Yes or No if your samples are purified. Typically enter NO, unless you gel purified your samples
9. Enter sequencing direction, number of samples to be sequenced (# of PCR products), total number of reads (# of PCR products x # of sequencing directions)
10. Click Next
11. Download template (in Primer Information section) and rename the file with your project name
- a. Under the Sample Information section: enter plate name, amount (20μL), and amplicon size (other columns can be left blank)
- b. Under the Primer Information section: enter plate name (fill out a row for each primer plate), concentration (3μM), and amount (25μL) (other columns can be left blank)
- c. Save file
12. Return to Beckman page and click Attach file to upload Primer File
13. Click Next
14. Enter date shipped, carrier (FedEx), and tracking # (if you have already generated FedEx slip, not required information)
15. Check .ab1 file
16. Shipping information should automatically be shown
17. Get quote and PO number (on top of freezer under thermocycler sign up board)
18. Submit Project and print a copy of the submission form
19. Login to our FedEx account:
- See password sheet in computer room for login and password information
20. Find Beckman shipping profile, which is saved under My Shipping Profile
21. Click Ship and Print
22. Package plates in separate ziplock baggies for each plate. Wrap with bubble wrap. Put inside Styrofoam box within a cardboard box
23. Package samples on dry ice (see Sarah for lab account usage)
- tape Styrofoam box closed
- include a copy of the PO and submission form in the cardboard box
- put FedEx slip on the top of cardboard box in provided sticky envelope
- make sure to put a dry ice form on the outside of cardboard box (diamond sticker)
Here is the shipping information just in case (standard overnight) to:
Beckman Coulter Genomics, Inc
36 Cherry Hill Drive
Danvers, MA 01923-2575
Fill out sign-up sheet on Post-PCR freezer (below thermocycler sign-up board).
- Very important for budgeting
Renaming .ab1 files
Beckman names the files according to the project name. The Shaffer lab provided us with a script to rename the files. Two files are required and can be found on the iMac named Brock3 in the seq renamer Beckman file (desktop).
1. Login to Beckman
2. Click on Get Your Results Tab
3. Download your project and save
4. Create an excel or text file with the names of your samples in one column.
- …through H1
- …through H2
- etc
5. Open new_names.tab
6. Change the first column to your project name with Find and Replace.
- Example:
- new_names.tab file = GGM9_A01 etc
- my project name = ss
- Find: GGM9
- Replace: ss
- do not change _A01
7. In your excel or text file, copy your column of sample names
8. Paste over second column in the new_names.tab file (in TextWrangler hold down control and drag curser over second column to highlight only the second column)
9. Save file
10. Copy and paste the new_names.tab and chromat_renamer.perl into the file that contains your .ab1 files
11. Open Terminal
12. Change directory to your file that contains your .ab1 files
- Type cd (drag your file into terminal window); press enter
13. Type perl chromat_renamer.perl; press enter
14. A new file named renamed_chromats should be in your .ab1 file with your samples renamed.
Ethanol precipitation for DNA clean-up
1. In the 1.5mL epi tube containing the DNA sample to clean, add 1mL of COLD 100% ethanol and 100ul sodium acetate (NaOAc) pH 5.2. Mix by inverting.
2. Place in -20°C freezer for at least 3 hours, overnight is fine.
3. Spin at 14,000 rpm in a chilled centrifuge at 4°C for 10-15 min.
4. Discard supernatant. Careful not to disturb pellet! Pellet may be invisible.
5. Re-suspend in 1mL fresh 70% ethanol, centrifuge for 5min at 14,000 rpm.
6. Discard supernatant.
7. Dry pellet in Speed Vac for up to 10min at low or medium heat.
8. Re-suspend pellet in 1x TE or Qiagen AE buffer. Generally use 50-200ul, depending on amount of DNA present. Thermomixer is probably ideal at RT.
Phenol/Chloroform extraction of genomic DNA
Day 1: Digestion of Sample
1. Finely mince ~25 mg of tissue or 200 µl of blood to a 2.0 ml tube.
2. Add the following reagents to the tube:
- 500 µl of 1x TEN Buffer
- 50 µl of 10% SDS
- 40 µl of Proteinase K (20 mg/ml)
3. Mix by vortexing
4. Incubate at 56 °C overnight on a rocking platform.
Day 2: PCI Extraction
- Make sure the relevant buffers (PCI, CI) and reagents (NaOAc, 70% EtOH) are prepared!
1. Remove tubes from incubator, and add 500 µl of buffered phenol. Shake tubes and invert several times (5 – 10 mins). Spin tubes in centrifuge at max speed (14,000 rpms) for 5 mins.
2. Observe 2 layers. The top layer contains the DNA/RNA and the bottom, organic layer the waste.
3. Remove the top layer and transfer into a new and labeled tube, and discard the bottom layer in the PCI organic waste container.
4. Add 500 µl of PCI to the samples in the new tubes, and shake tubes and invert several times (5 – 10 mins). Spin tubes in centrifuge at max speed (14,000 rpms) for 5 mins.
5. Observe 2 layers. The top layer contains the DNA/RNA and the bottom, organic layer the waste.
6. Remove the top layer and transfer into a new and labeled tube, and discard the bottom layer in the PCI organic waste container.
7. Add 500 µl of CI to these new tubes, and shake tubes and invert several times (5 – 10 mins). Spin tubes in centrifuge at max speed (14,000 rpms) for 5 mins.
8. Pipette off the top layer and transfer it to a 1.7ml final epi tube with the final sample name and number.
9. [OPTIONAL] If you want to RNase treat the DNA, add 1ul of a 10 µg/mL stock solution of RNase A to your DNA and incubate at 37ºC for 30-60 min.
10. To this tube add the following:
- 100 µl of 3M sodium acetate (NaOAc)
- 1 ml of cold 100% ethanol (EtOH)
11. Invert tubes several times and place in a -20 freezer overnight. (3 hours is sufficient).
12. You can either stop here and do precipitation the next day or after 3 hours, complete the precipitation.
Ethanol Precipitation
1. Remove tubes from the freezer, and centrifuge at max (14,000 rpms for ~10 min) to pellet the DNA.
2. Observe a white or brownish pellet. (May be absent in low concentration samples).
3. Decant the supernatant, being careful not to lose the pellet.
4. Add 1ml of fresh 70% EtOH to the tube containing the pellet. Vortex briefly to re-suspend the pellet.
5. Centrifuge at max (14,000 rpms for 10 min) to pellet the DNA.
6. Decant the EtOH, being careful not to lose the pellet.
7. Remove the remaining 70% EtOH by vacuum centrifuging in the tubes in the Savant Speed-Vac for 10 mins or until the pellet is dry. (Do not use high heat for more than 5 mins).
8. Re-suspend DNA in 100-200 1x TE (or Qiagen's AE) Buffer.
RNase treatment [if you didn't do this prior to precipitation]
- If you are extracting DNA from an RNAbuffer (e.g. PAXgene RNA tubes which do not contain any DNases), you may consider doing an RNAse treatment of the final DNA*
1. Add 1ul of a 10 µg/mL stock solution of RNase A to your DNA and incubate at 37ºC for 30 min.
2. Recover the DNA by adding 1/10 volume of 3M sodium acetate (pH 6.8) and 2 volumes of isopropanol or 95% ethanol to the DNA containing solution.
3. Incubate on ice for 10 min.
4. Centrifuge at maximum speed for 5 min at room temperature, to pellet the DNA.
5. Discard (carefully) the alcohol. Wash with 70% ethanol and dry DNA via Speed-Vac on low heat for 10 mins.
6. Dissolve in AE, TE or dH2O, as you did in the DNA extraction.
Common buffers and preparations by the final desired volume:
TEN Buffer 1x | 50mL | 25mL |
0.5M EDTA | 100μL | 50μL |
5M NaCl | 1mL | 500μL |
1M TRIS | 500μL | 250μL |
ddH2O | 48.41mL | 24.2mL |
PCI | 50mL | 25mL | 12.5mL |
Buffered Phenol | 25mL | 12.5mL | 6.25mL |
Chloroform | 24mL | 12mL | 6mL |
Iso-Amyl Alcohol | 1mL | 500μL | 250μL |
CI | 50mL | 25mL |
Chloroform | 48mL | 14mL |
Iso-Amyl Alcohol | 2mL | 1mL |
70% Ethanol | 50mL |
100% Ethanol | 35mL |
ddH2O | 15mL |
Calibration of SeraPure beads to AMPure XP beads
Note: This protocol is derived via Brant Faircloth from the referenced protocol created by Nadin Rohland (Rohland N, Reich D. Cost-effective, high-throughput DNA sequencing libraries for multiplexed target capture. Genome Research. Early Online Access. Doi: 10.1101/gr.128124.111)
1. Prepare fresh aliquots of 70% EtOH (70ml EtOH + 30ml water).
2. Mix 1μg ladder per trial volume.
- Use 20μL 50bp E-Gel ladder (undiluted).
3. Add trial volume of serapure to 20μL of ladder (may need to do more trials if Serapure haven't been tested in awhile).
- Example:
Final concentration of Serapure | Volume of ladder | Volume of Serapure |
.2X | 20μL | 4μL |
.4X | 20μL | 8μL |
.6X | 20μL | 12μL |
.8X | 20μL | 16μL |
4. Mix gently with a pipette and incubate at room temperature for 5 minutes.
5. Place tubes in magnetic tube holder and allow to rest for 2 minutes. Beads should be drawn out of solution and should gather on the side of the tube in contact with the magnet.
6. Carefully pipette off supernatant - without disturbing the beads - and discard.
7. Add 500uL fresh 70% ethanol to each tube, then pipette off ethanol and discard.
8. Repeat step 6.
9. Allow beads to air dry with tube caps open to remove all traces of ethanol (~5-10 minutes) or place beads of a 37°C heat block for 3-4 min until dry.
10. Resuspend beads in 10μLuL 1X TE buffer to elute DNA.
11. Transfer elute to new labeled tubes and discard beads.
12. Mix elute with 1μL of loading dye.
13. Electrophorese in an E-gel for approximately 9 min.
14. Compare the results of the trial volumes do obtain the most appropriate Xul needed to do the following AMPure Bead Cleanup Protocol or for library preps.
15. Post results on small Post-PCR refrigerator.
AMPure XP bead clean-up of DNA
1. Use sample DNA concentration to calculate volume that will yield 1μg of DNA.
- To calculate the number of microliters needed to obtain 1μg of DNA, divide 1000 by the concentration of DNA in ng/μL.
- To obtain genomic DNA, use a ratio of XuL bead solution per μg of sample DNA (X is the optimum volume figured out by testing the ampure mixture; typically 40-60μL).
2. Obtain clear 1.7mL tubes. In each 1.7mL tube, combine bead solution, DNA, and 1X TE buffer in the following ratio: XμL : 1μg : up to 160μL total volume.
- Can be scaled up to combine as much as 3μg DNA with 3XμL bead solution and 1X TE to a total volume of 480μL in each tube.
3. Mix gently with a pipette and incubate at room temperature for 5 minutes.
4. Place tubes in magnetic tube holder and allow to rest for 2 minutes. Beads should be drawn out of solution and should gather on the side of the tube in contact with the magnet.
5. Carefully pipette off supernatant - without disturbing the beads - and discard.
6. Add 500μL fresh 70% ethanol to each tube, then pipette off ethanol and discard.
7. Repeat step 6.
8. Allow beads to air dry with tube caps open to remove all traces of ethanol (~5-10 minutes) or place beads of a 37°C heat block for 3-4 min until dry. You can also use a sterile toothpick to remove any blobs of ethanol from the tube walls.
9. Resuspend beads in 10μL 1X TE buffer to elute DNA.
10. Transfer elute to new labeled tubes (combining duplicates from the same initial sample, if possible) and discard beads.
11. NanoDrop to determine new sample concentration. Perform ethanol precipitation to concentrate if necessary.
Ambion Trizol Plus RNA purification kit protocol
1. Turn centrifuge on and set to 4°C.
2. Clean area with bleach/RNase away. Set up mortar and pestle with tinfoil, petri dish, and razor blade.
3. Label round-bottomed tubes.
4. Add 300ul TRIzol to tubes.
5. Put samples on ice.
6. Using tinfoil covered mortar and pestle, crush the vial containing the sample.
7. (optional: on dry ice) Using the razor blade cut the sample into pieces and quickly transfer into TRIzol.
8. Homogenize samples for 3min using bead mill.
9. Add 700ul TRIzol.
10. Incubate at room temperature for 5 minutes.
11. Add 200 μl chloroform and shake the tube vigorously by hand for 15 seconds.
12. Incubate at room temperature for 2-3 minutes.
13. Centrifuge the sample at 12,000 x g for 15 minutes at 4°C.
- Make up 1ml of 70% ethanol for each sample.
- Make up Purelink DNase mixture (10ul DNase, 8ul buffer, 62ul water).
14. Transfer ~600 μl of the colorless, upper phase containing the RNA to a fresh RNase-free tube.
15. Add an equal volume of 70% ethanol.
16. Mix well by vortexing and invert the tube to disperse any visible precipitate.
17. Transfer up to 700 μl of sample to a spin cartridge (with a collection tube).
18. Centrifuge at 12,000 x g for 15 seconds at RT. Discard the flow-through and reinsert the spin cartridge into the same collection tube.
19. Repeat steps 17-18 until the entire sample has been processed.
20. Add 350 μl Wash Buffer I to the spin cartridge containing the bound RNA. Centrifuge at 12,000 x g for 15 seconds at RT. Discard the flow-through and collection tube. Insert the spin cartridge into a new collection tube.
21. Add 80 μl Purelink DNase mixture directly onto the surface of the spin cartridge membrane.
22. Incubate at RT for 15 minutes.
23. Add 350 μl Wash Buffer I to the spin cartridge. Centrifuge at 12,000 x g for 15 seconds at RT. Discard flow-through and collection tube. Insert the spin cartridge into a new collection tube.
24. Add 500 μl Wash Buffer II with ethanol to the spin cartridge.
25. Centrifuge at 12,000 x g for 15 seconds at RT. Discard flow-through and reinsert the spin cartridge into the same collection tube.
26. Repeat steps 24-25 once.
27. Centrifuge the spin cartridge at 12,000 x g for 1 minute to dry the membrane with bound RNA. Discard collection tube and insert the spin cartridge into a recovery tube.
28. Add 30 μl RNase-free water to the center of the spin cartridge.
29. Incubate at room temperature for 1 minute.
30. Centrifuge spin cartridge and recovery tube for 1 minute at ≥12,000 x g at room temp.
31. Insert the spin cartridge into a new, labeled recovery tube.
32. Add 20 μl RNase-free water to the center of the spin cartridge and repeat steps 29-30.
33. Place on ice. Take 3ul for nano drop and surface tension gel.
34. Store at -80°C as soon as possible.
Obtaining DNA from degraded samples for NGS sequencing


text:
Neo FreeRunner Hardware Issues
From Openmoko
Neo FreeRunner
This is a community-written page that discusses hardware issues with the FreeRunner/GTA02 device. Information here is unofficial (and possibly incorrect) unless otherwise stated. Corrections and clarifications from Openmoko employees would be greatly appreciated.
Please DON'T PANIC when reading this page. Please give Openmoko employees time to investigate these issues and to develop a solution. Some of the items may turn out to be non-issues, or may have software workarounds. The FreeRunner's software is still under heavily development and can help fix most of this problems.
Please always file a bug report on the issues and mention the bug report number in this page. Otherwise it is impossible to check if a bug has been fixed or not.
Contents |
Active Issues
Poor Audio Quality (FIXED)
Please use http://docs.openmoko.org/trac/attachment/ticket/2121/gsmhandset.state.new for *all* Neo Freerunner. (2009-04-27 jOERG)
Issue: The person on the other end of a GSM phone call may experience poor audio quality, to the point where he/she cannot carry on a normal conversation.
There are three related aspects to this issue:
- Call is too quiet (mixer settings for mic and/or earpiece are set too low)
- Caller hears a loud echo of their own voice #1267
- Buzzing noise caused by GSM radio interference #883 #1352 See GSM buzz.
Affects: all devices but only some users (depending on many factors)
The source of the GSM buzz has been identified. It's a mere hw-issue, depending on the way you hold the device, and the local situation created by network settings made by the GSM-provider (mainly 1800/1900 seems to be affected), as well as your position relative to basestation. There is no way to fix GSM buzz by mixer-setting modifications. So all suggested settings here may improve a little the volume of buzz during you're *not* speaking only, while relative ratio of buzz/voice level while speaking can't be changed by mixer settings.
Some hidden Calypso commands may help with the echo problem.
hexedit, ghex /opt/Qtopia/plugins/phonevendors/libficgta01vendor.so search "AT%N0125" and change to "AT%N0187"
- Using a bluetooth headset is a possibility.
- Using external GSM-antenna will stop buzz.
Due to the multiple factors influencing the result of a single test, it's nearly impossible to find a setup that lets you compare for a decent "before / after" result. If you ever took an old analog TV portable to a place where you had to fiddle around with the antenna to try and make the snow and shadows vanish off the picture, you might have gotten a slight idea of what it's like to reproduce the same situation for decent tests. So probably most of the suggested alsa-improvements are mere random results. Even if they worked for provider A evidently, this doesn't mean there's any improvement by using same settings for provider B.
This being said, here they are:
- Better mixer settings: One confirmed good settings are here: http://email@example.com/msg00564.html. Please change accordingly in /usr/share/openmoko/scenarios/gsmhandset.state. Should eliminate/lessen echo and buzz problems. Neo alsamixer is the main article for setting the mixer settings.
Better set of mixer settings
Regarding 2. in the previous paragraph, here are my further tweaked settings:
[mic volume & buzz problem]
- 'Mono Playback Volume' (95)
- 'Mono Sidetone Playback Volume' (2)
- 'Mic2 Capture Volume' (3)
[speaker volume & echo problem]
- 'Speaker Playback Volume' (112)
- 'Bypass Playback Volume' (5)
Lowering Mono Sidetone eliminated the buzz problem better. Please try out and report if you have good success... --TimoJyrinki 08:19, 16 October 2008 (UTC)
Can't boot with discharged or missing battery (FIXED)
Issue: Neo FreeRunner requires battery power to boot, because Neo FreeRunner consumes too much current while booting to boot with only a charger. Since charging isn't enabled until the Neo FreeRunner has booted, this means that a discharged battery can not be charged. Some versions of uBoot don't continue booting and just charge bat (red LED flashes constantly) when probing bat voltage shows it's too weak to start FR. Let the device sit and charge for an hour, then reboot.
Affects: All A5, older A6. With change of Vsys buffer capacitor to 100uF this issue has been fixed for good.
Fixed(?) with Werner Almesberger patches - more here additional information
I think this patch finally fixes the problem: Philipp Hugs patches
- First test if your FR starts with no bat inserted. If it does you can forget about this whole topic.
- Make sure that the battery never discharges completely
- Use external stand-alone charger (compatible with the Nokia BL-5C battery)
- Boot the FreeRunner with an alternative battery, or with a spare GTA01 or GTA02 battery, plug USB power, then switch to the empty battery.
- Boot the FreeRunner with a 4.5VDC external power source (steady hand and great care involved), plug USB power, then insert the empty battery.
- Oscilloscope digital clip probes fit perfectly on the Freerunner's battery connectors. Make sure polarity is correct!
- Some users have reported that Neo FreeRunner is able to boot on USB power alone using the NOR u-boot, thus: press AUX, plug USB power, select boot. Reports of success would be appreciated.
- Does work with sequence described. If after pressing boot it just turns off again (happened a few times), then: press AUX, plug in USB power, cycle through the menu a few times to keep it from turning off without booting, after a minute or so press boot. Imrehg 08:34, 10 September 2008 (UTC)
- This does indeed work with sequence described. After pressing boot, the screen went blank for a few seconds, but then the beast came to life again. That saved the day! Edictor
- Trifirmed. Neither USB connection nor wall charger was able to wake my phone. But in NOR u-boot it worked (connected to the computer). I think this trick hasn't failed to anyone. --Flamma 12:11, 17 September 2008 (UTC)
- Unfortunately this is not working for me. My FreeRunner has been off for a few weeks and completely dead. I tried various combinations of holding aux and plugging in to both wall and computer, no luck. Tried with and without the battery in place, no luck. Aux and power button / just power / aux then power. No luck... --Safire 6 November 2008
- This is working ! It has prevent me from buying an external power charger. My freerunner had its battery discharged for about 2 weeks now, and I was able to boot and charge it. --JRD 23:37, 12 November 2008 (UTC)
- It has worked for me ! I have booted to 2008.9 Om with discharged battery (after connecting the FR to power supply and waiting a few minutes until AUX button red light stopped flashing) using the NOR boot menu. Tried the same with NAND boot menu and worked also !!! Then I could charge the battery. -- Emsyr 3:23, 20 November 2008 (UTC)
- I can boot to 2008.9 without battery and with USB connection = plug USB power (FR <-> PC) + press power button. However I cannot shutdown correctly. --Cynan 13:43, 9 December 2008 (UTC)
- I can boot to 2008.9 without battery and with USB connection: press AUX button, then plug USB power, then press power button (keeping AUX pressed). NOTE that battery was fully charged. The problem wasn't battery but corrupted environment in NAND u-boot: with battery IN, no access to NOR u-boot
- Both my friend's freerunner and mine was completely discharged. This trick failed. Finally booted the freerunner with a BL-5C battery from an old nokia n-gage. With the power connected I then switched the batteries. --Unlotto 14:27, 25 January 2009 (UTC)
- Another confirmation that this works. You may use the computer-USB cable or the AC adapter. I held AUX, plugged in the USB, and the NOR boot menu started before I even hit the Power button. For some reason, I have to cycle through the menu once before I choose boot, otherwise it will power off as soon as it loads the kernel. I booted it, let it start up fully, and it continued charging. After a minute or two, as a test I shut down and tried rebooting normally. There was enough power in the battery at that point to boot normally from the AC adapter. Note, I did not encounter this problem (using the AC adapter) until after I "upgraded" from the pre-installed May 2008 u-boot to the December 2008 u-boot; perhaps that is relevant. --Robolange 05:53, 1 February 2008 (UTC)
- This did not work for me (with the 1.3.2-moko12 u-boot). However, I was able to "jumpstart" my Freerunner with a bench supply and some micro-clips, as suggested above. (Worried I would fry something, I tried at lower voltages first, but it didn't start up until I set the supply to 4.5v, with the OM USB charger also attached.) Once it was well into the Linux kernel boot sequence I removed the clips and put in the battery to charge. --Wiml 05:30, 9 April 2009 (UTC)
- This worked for me. Now I'm not afraid of letting my FreeRunner discharge. Cristianpark 20:00, 30 August 2009 (GMT -5)
See also:
Suspend/resume corrupts SD card's partition table
Issue: Suspend/resume corrupts the partition table of the SD card
Affects: Unknown.
[Patched kernel] might solve the problem
- [Script] as a temporary workaround
See also:
Known/Accepted Issues
This section lists items that are acknowledged as being less than ideal, but are considered to be acceptable in the shipping product. They will not be discussed in detail on this page.
- Poor performance + slow bus speed of the Glamo GPU - discussed to death on the mailing lists and IRC.
- stable-2.6.26 branch of kernel has wait states lowered, core speed increased from 50MHz to 80MHz and memory speed from 80MHz to 90MHz (the latter is also in stable branch) - these lessen the problem a bit, though it's slow still
- GPS antenna switch driven out-of-spec - does not appear to have a significant effect on device performance
- Poor low-frequency audio response with low-impedance headphones, e.g. as discussed in this thread: http://lists.openmoko.org/pipermail/openmoko-kernel/2008-March/001999.html (NOTE - this thread refers to pre-mass-production devices)
- Can be fixed to an semi-acceptable level (if not high fidelity most probably) by adjusting "Bass Volume" to full (15) and "Bass Filter" to "100Hz @ 8kHz" (bass will be boosted <= 600Hz when playing back at 48kHz) or "200Hz @ 8kHz" (<= 1200Hz @ 48kHz). The default is 130Hz @ 48kHz and does not help much with the more wider scope of low frequencies.
- Ideally someone would record output and find out which setting produces best output, ie. compensating for the loss of low frequencies without boosting too high frequencies with this "bass" boost.
- There exists a hardware fix.
Actually with 16 Ohm headphones the cutoff frequency is more than 2kHz, so even 1200Hz @ 48kHz seems to be not appropriate. If you have 30 Ohm, it might be just correct setting.
Resolved Issues
These are issues that have been discussed in the past, but have been fixed (or turned out not to be a problem) for the mass-produced devices.
- Excessive LED current - Some early units lacked a current-limiting resistor for the LEDs. This has been fixed for the production units.
- Battery life - At this time it appears that the FreeRunner battery life will be acceptable once suspend/resume support has been implemented in software.
- slow GPS TTFF - see GPS Problems
Battery discharges when charging completes
Issue: If the Neo FreeRunner has been charging, when charging completes, it seems to drain the battery and not turn on charging again. This seems to be bug of PMU-registers setup, that shows up when PMU has to handle bat autonomously (=suspend). There might be issues our current scheme relies on wake-interrupt at bat-full which doesn't succeed, or something like that.
Affects: Unknown.
This has been fixed in linux. See http://docs.openmoko.org/trac/ticket/1158
Some SIMs Don't Work
See FreeRunner unable to work with 3G SIM cards
Status: Fixed in GSM firmware moko10-beta2 or later. See GSM/Flashing for instructions.
Empty NOR Flash
Issue: NOR flash (backup copy of u-boot) is not programmed. #1568
Affects: Unknown - maybe only 1 or 2 devices? Also need to confirm that the bug report was from a mass-production unit rather than an earlier prototype.
Status: Solved. The new devices are tested better if they have the NOR flashed programmed and aren't shipped if the test fail.
Workarounds: Use the NAND copy of u-boot and be careful not to brick the device unless a debug-board is available.
Debug board has wrong vendor/product ID
Issue: A debug board is not reporting the expected USB Vendor/Product ID.
Affects: Only one report has been seen so far.
Status: Probably a single case scenario.
Workarounds: Edit configuration files (e.g. openocd.conf) to use the IDs that the board is reporting.
Possible hardware fixes on GTA02
List of "Current issues" Imported from the "Community update page"
(to be sorted)
The information below has been collected from various sources, feel free to add questions and comments here.
GPS Performance of the FreeRunner
The poor GPS performance on the FreeRunner has been traced to an interaction between the microSD card and the GPS unit. A software and a hardware fixes are available, see GPS Problems.
GTA02 battery status
While writing a device driver for the new battery which provides an accurate counter of the charge state of the GTA02, the driver developer discovered that the device driver does not get a reading of the charge state due to a very long response time with only one I/O signal when trying to read the charge state. To be able to read the battery status properly, it has been written that it will be necessary to re-design that part of the GTA02 for hardware version GTA02A5 to use two I/O signals to reduce the response time (one for transmitting commands, one for receiving data?). This was fixed (see Bug 957).
SMedia 3362 Documentation & OpenGL ES Drivers
There is an open source kdrive driver being written for the GTA02 which will use hardware to accelerate the XRender extension. While the chip is capable of 3D graphics, no OpenGL ES driver/library is avaliable and Openmoko developers will not be writing one in the near future (although they have not ruled it out in the long-term).
Documentation for the SMedia 3362 has been promised (see this post). However, this refers to documentation Openmoko developers will be writing themselves, not the technical documentation SMedia have provided Openmoko with. The Openmoko developers had to sign an NDA with SMedia to obtain this documentation and are therefore unable to pass this information on to community developers. (See this post for details)
Draws too much current from USB
It may be that the Neo draws too much current from your USB host/hub, and that the USB host/hub switches off to prevent damages.
The behaviour is controlled by a complex interaction between soft- and hardware, see Forcing fast charge mode and USB host. It may be that the host negotiates more power than the Hub is willing to provide. In case of problems, try to limit the current to e.g. 400mA.

subset: commoncrawl


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You are watching : Camille Juban on Tahiti
Tahiti gets your heart pumping. I had been to Tahiti 4 times before. Sailed all over the island including Teahupoo. The island is amazing, the people show what the word Aloha should mean, but the water, being in this magical water is what sucks you in to this magical place. Windsurfing at Teahupoo is something you can never understand. Watching these kids charge it without any fear inspires me like never before. Filming Camille drop in to a barreling mast high Teahupoo wave, set up and hit the lip over this wave is indescribable. His speed, timing, and commitment is unbelievable. As I sat on the boat watching, filming, trying to grasp what was going on, the fear in me boiled as I waited for my turn. While we didn't have anyone to film when I went out, it was ok. It quickly made me realize how I wanted to show the images of Titun and Camille charging this place. Suddenly I wanted to swim in and capture more of the moment, more of the feeling, try to get the audience to feel and see the power of this place. The first reason I ever picked up a video camera was to try and show the world what windsurfing was from the inside. To capture it like a windsurfer sees it. I hope this video makes you feel the passion of three guys traveling to one of the most epic windsurfing places on earth, shredding waves, taking turns, having fun, and being blessed with one of the most memorable windsurfing days of their career. Music: ALT-J "Fitzpleasure"
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Construction Equipment Rental in Reelsville, Indiana
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There is little or no wait time involved before you can start listing out your construction equipment rental needs. We will make sure the vendors get in touch with you on the same day.
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Dial our toll free number today. Let us make the arrangements for you. You can also submit a short contact form on the right to get in touch with us.
Equipment Rental Reelsville, Construction equipment rental Reelsville, Construction Reelsville,Heavy Equipment Rental Reelsville.


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Mr. Brown is hoping that Washington will pony up more than $50 billion, but the feds have committed only $3.3 billion so far—and Republicans intend to claw it back if they take the Senate and White House this fall. If that happens, the state won't have enough money to complete its first 130-mile segment in the lightly populated Central Valley, which in any event wouldn't be operable since the state can't afford to electrify the tracks....What a disaster! Remember a year and a half ago, when Wisconsin elected a new governor who said he'd kill the high-speed rail deal? I was a single-issue voter for Scott Walker then, and that was my issue. Here's a newspaper article from December 2010: "High-speed rail funds scatter to other states... California gets lion's share."
Wisconsin will keep only a fraction of the $810 million it won in federal high-speed rail money.... California is the big winner, with up to $624 million....Ha. It's been so not tragic.
[Exiting Democratic Governor Jim] Doyle called the loss of the high-speed rail funds a "tragic moment for the state of Wisconsin. Eight hundred and ten million dollars that would have gone to create thousands of jobs in Wisconsin will now create jobs in other states... I obviously am deeply saddened to see us take a major step backward."
"My congratulations to the workers in California and Florida. As a result of this decision, you will have a merry Christmas," Milwaukee Mayor Tom Barrett said at a news conference. "I'm just sad the same won't happen here in Wisconsin."Tom Barrett is, of course, the Democratic Party candidate in the recall election against Scott Walker, but back then, he'd just lost the regular election to Walker. The recall election is in 2 weeks. I wonder if he's been asked about these old statements about the damned train and how lucky California was to get that federal cash infusion Scott Walker spurned.


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Do you remember those cool fall nights under the lights at the football field? No matter what crowd you hung out with, we all remember attending the games!
Take a trip back in time this season and go out to cheer on the team. Invite some of your old classmates to join you in the bleachers and bundle up in some fan gear to show your true school spirit!
Go Patriots!
If this e-mail doesn't load properly, click here to view online. To stop receiving these e-mails, just click here. birmingham.alumniclass.com has no association with this school or school district.


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TV on the Radio
TV on the Radio's story is one of synchronicity, of serendipity, of epiphany. TVOTR began with Tunde Adebimpe (vocals) and David Sitek (multi-instrumentalist and production kingpin behind the Grammy nominated Yeah Yeah Yeahs LP Fever To Tell) being roommates, drinking way too much coffee, making art: comics, movies, paintings. They started trading tapes of each other's four track recordings and thought it'd be fun to try to make "a real record" out of all that stuff.
Touch & Go released TV on the Radios stunning EP Young Liars in July 2003, and it immediately marked them out as one of the most musically innovative bands to have emerged in years.
Their debut full-length Desperate Youth, Bloodthirsty Babes set 2004 off to a dramatically positive start. With new member Kyp Malone sharing songwriting, guitar and vocal duties, TVOTR was quickly on the path to the being one of the most talked-about new groups in America. After touring as a trio, Jaleel Bunton (drums) and Gerard Smith (bass) joined the band officially.
TVOTR has gone on to enjoy critical acclaim in esteemed publications ranging from The New York Times to SPIN Magazine and from Rolling Stone to URB, and performed live on The Late Show With Carson Daly and the Late Late Show. NPR has featured them on several programs, most notably All Things Considered.
TV on the Radio capped a year of critical acclaim and non-stop activity when their debut album Desperate Youth, Bloodthirsty Babes was awarded the Shortlist Music Prize in Los Angeles on November 15th, 2004. The Shortlist Prize was founded in 2001 "to honor the most creative and adventurous albums of the year across all genres of music".


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