Team:Penn/Notebook

University of Pennsylvania iGEM

2015 iGEM Lab Protocols

iGEM, University of Pennsylvania
Lab: 220s South 33rd Street, 230S Skirkanich Hall
Philadelphia, PA 19104-6321

Colony PCR


Protocol: Template Preparation 

  1. First, one must have a monoclonal E Coli colony source. This may be one of the following three things:
    1. A glycerol stock made from a SINGLE colony on an agar plate
    2. An overnight culture in LB media (+ antibiotics if appropriate) started from a SINGLE colony on an agar plate
    3. A SINGLE colony on an agar plate
  2. Label a sterile eppendorf with the strain number and assign a unique colony number for each colony picked. Pipet 20 uL of sterile water into the tube. If the e coli is coming from sources (1) or (3), use a clean, sterile pipet tip to scrape a tiny bit of bacteria (even the very smallest amount is okay) and then pipet up and down into the waiting 20 uL. If the e coli is coming from source (2), pipet 1 uL of the LB culture into the water. Mix well.
  3. This water/colony mixture will be the “template” for your PCR reaction. Next, set up your reaction mixture and run the cycler protocol. Follow this up by running a gel to examine the PCR amplicon lengths.


Reaction Preparation 

  1. 1 uL Template
  2. 10 uL 2x Taq Master Mix (Long term storage in freezer, will last one month at 4C)
  3. 1 uL FWD Primer (@ 10 uM)
  4. 1 uL REV Primer (@ 10 uM)
  5. 7 uL H2O


Cycler Protocol- The cycler protocol is fixed except for the extension time. Set the extension time for 1 min per kb of desired amplicon at 68C, plus a little bit of buffer time. For example, for a 2 kb amplicon, set the extension time to 2 min 30 seconds.

  1. 95ºC for 6 minutes
  2. 30x [95ºC for 30 sec, 55ºC for 30 sec, 68ºC for 1 min/kb amplicon]
  3. 68ºC for 20:00 min
  4. Hold at 4ºC

DNA Digestion


Reaction Mixture Preparation

  1. 5 uL 10X Digest Buffer
  2. 1 uL of each digestion enzyme up to maximum of 2.5 uL
  3. 2000 ng of desired cut fragment up to max volume of 50 - enzyme volume (cutsmart+digestion enzyme volumes)
  4. Fill with milliQ water to 50 ul


Digestion Protocol 

  1. Incubate the reaction at 37ºC for 1 hour.
  2. The digest buffer is specific to the enzyme and it stated on the enzyme product description - look it up on NEB’s webpage for each enzyme. It is generally “Cutsmart.” For digests with multiple enzymes, use NEB’s tool to identify the right buffer.
  3. Enzymes are stored in glycerol, but glycerol can inhibit digestion reactions. To keep the volume to 2.5 uL, use 1 uL of 2 enzymes. For three enzymes, consider using just 0.5 uL of the most active one or reducing the volume of all three to 0.8 uL.
  4. You want there to be at least 2000 ng of the fragment you want to isolate in the end. This is because DNA digestion products must be purified, a process that is only about 50% efficient. Isolating 2000 ng ensures you have enough DNA in the end for cloning. To calculate the volume, use the DNA concentration and the relative fraction of the plasmid that is the fragment you want. For example: for a 10 kb plasmid at 1000 ng/uL with a 1kb fragment I want to cut out, I’d want to do the following calculation:

DNA Gel Extraction


Gel extraction: Follow the Qiagen QiaQuick DNA Gel Extraction protocol that accompanies their kits. Below are the essential steps:

  1. Add 3 volumes of QG buffer per 100 mg of agarose gel (i.e. add 450 uL of QG buffer for a 150 mg agarose gel chunk)
  2. Heat at 50ºC for ~10 minutes until the gel completely dissolves. Vortex to mix.
  3. Add 1 volume of isopropanol per 100 mg of agarose gel (i.e. add 150 uL of Isopropanol to a 150 mg agarose chunk dissolved in 450 uL of QG buffer)
  4. Vortex to mix.
  5. Add up to 700 uL isopropanol-QG-DNA-Agarose mixture to a gel extraction column (purple with lids). Spin down at 13000 rpm for 1 minute. Discard the flow through.
  6. Repeat step 5 until you have spun down all the dissolved gel you have
  7. Pipet 750 uL of PE buffer (**Make sure 200 proof Ethanol has been added) onto the column
  8. Spin down at 13000 rpm for 1 min. Discard the flow through.
  9. Air dry by spinning down at 13000 rpm for 1 min.
  10. Move the column to a fresh eppendorf tube. Pipet 35 uL of EB buffer above the column’s membrane - careful not to puncture it!
  11. Let the EB buffer sit on the membrane for 1 min.
  12. Spin at 13000 rpm for 1 min to elute the purified DNA into the eppendorf tube.
  13. Consider measuring concentration on the nanodrop (see separate protocol)


 

DNA Purification


It is best to follow the QiaQuick PCR Purification protocol supplied with their kit. Below are the essential steps: (All buffers and columns are supplied with the QiaQuick PCR purification kit.)

  1. Add 5 volumes of PB (binding) buffer to the DNA sample you’d like to purify. For example, if you have a 50 uL completed PCR reaction to cleanup, start by adding 250 uL of PB buffer and mixing well.
  2. Add the PB + DNA mixture to a PCR cleanup column (purple columns, same as gel extraction). Spin at 13000 rpm for 1 min. Discard the flow through.
  3. Pipet 750 uL of PE buffer onto the column. Spin at 13000 rpm for 1 min. Discard the flow through.
  4. Air dry the column by spinning at 13000 rpm for 1 min. Transfer the dried column to a fresh eppendorf tube.
  5. Add 35 uL of EB buffer to the column and allow it to stand for 1 min.
  6. To elute, spin at 13000 rpm for 1 min. The solution that flows into the eppendorf collection tube is your purified DNA. Consider measuring its concentration with the nanodrop.

Gel Electrophoresis

  1. Unwrap a new E-Gel and snap it into the E-Gel Holder. Make sure the gel holder lights up to indicate that the E-Gel is properly connected.
  2. For the 1.2% 12 well E-Gels, add 15 uL of H2O to each well that will receive a sample, and add 5 uL of sample (for a total of 20 uL). You may load all 20 uL for a colony PCR reaction to ensure bright bands for even the smallest of amplicons. NO loading buffer is required - do not add any!
  3. Add 20 uL of 2-log DNA Ladder (at 1 ug/20 uL) to one lane
  4. Run for 15 minutes-30 minutes and visualize on the system itself (SAFETY: Only visualize the gel when the orange protective cover is on! Do not look directly into the blue light)
  5. Capture image on gel camera for long-term preservation of the result

 

Inoculating Cultures

  1. Obtain a sterile SNAP top culture tube and label it with the strain ID of the bacteria you’d like to grow
  2. Fill with 3-5 mL of LB (plus antibiotic if appropriate) using a sterile serological pipetter and sterile technique


If inoculating from a glycerol stock: Put a sterile pipete tip on a P200 pipetter as if you were to pipet a liquid. Touch the tip to the surface of the glycerol stock so a tiny bit of it is on the tip. Minimize time the glycerol stock is out of the -80ºC freezer to prevent freeze/thaw.


If inoculating from a single colony on an agar plate: Put a sterile pipete tip on a P200 pipetter as if you were to pipet a liquid. Touch the tip to the surface of the colony so a tiny bit of it is on the tip. I find holding the plate above a dark surface helps to visualize the colony and reduce glare. It is optimal to come at the plate with the tip from a roughly 45º angle and to just touch the surface of the colony without gouging the agar underneath.


Then for both glycerol stock/agar plate colony cultures:

  1. Pipet up and down with the bacteria-stabbed tip to mix.
  2. Replace Snap Cap top on culture top such that it is down a single notch (snap cap tops may be pushed to two stops. The first maintains the top on, but allows it to wiggle up and down. The second stop, achieved by pushing the cap further down, locks it into place). You want to have to cap on, but NOT in the locked position, such that the culture may be aerated.
  3. Place the tube in a shaking incubator set to 250 rpm and your growth temperature. For most E Coli work, 37ºC is appropriate for growing cultures. See the storage and experimental temperatures section for more detail.

Ligation


To calculate the volumes to be used, try this ligation calculator.

  1. 5 fmols backbone
  2. 30 fmols insert
  3. 1 uL 10x T4 ligation buffer
  4. 0.5 uL T4 ligase
  5. Fill to 10 uL with MilliQ Water


Mix everything together.*If you get a negative value for water, add 2 uL of ligase buffer and 1 uL ligase and make a 20 uL ligation reaction. Add 2 uL of ligation product for transformation 


Quick Ligation

  1. Incubate mix at 37ºC for 30 min
  2. Chill to 4ºC until ready to use 


Overnight Ligation

  1. Incubate mix at 16ºC overnight
  2. Chill to 4ºC until ready to use 

Making Media


LB Media

  1. Measure out 25 grams of LB broth powder and add it to a 1L vessel
  2. Fill to 1L with RO water
  3. Shake and allow to sit until most of the LB broth dissolves
  4. Loosen cap and mark with autoclave tape
  5. Autoclave on the liquid setting (“9” on floors 3,4 in Skirkanich)
  6. Tighten cap and allow to cool to room temperature before use


M9 Media (1L):

  1. 500ml 2xM9 salts 
  2. 30ml 10 mg/ml thiamine hydrochloride
  3. Dissolve 10 mg per ml of H2O
  4. Filter-sterilize using a 0.22μm filter
  5. Light-sensitive: store covered
  6. 10ml 40% glycerol 
  7. 20ml 10% casamino acids 
  8. 20ml 0.1M MgSO4
  9. 200μl 0.5M CaCl2
  10. 419.8ml sterile deionized H2O


Set up a sterile filter column and add each component separately into a 1 L bottle. Add antibiotic as appropriate and store at 4°C