7-24-14

added 250ul of chloramphenicol to LB

Put 4ml of LB+Chlor into 4 15ml eppindorf tubes

inoculated with colonies 3-3, 5-2, 5-3, and 5-4 using a wooden stick

put in 37oC on rotator overnight

On 10/2/14, we made a plate to make sure that the following samples would actually work:

• Red 1
• Red 2
• Orange 1
• Orange 2
• Cyan 1
• Cyan 2

Here are some useful links:

http://en.wikipedia.org/wiki/Green_fluorescent_protein

http://en.wikipedia.org/wiki/Fluorescence_spectroscopy

 

Orange 1 does not work.  All of the others look good.  See the attached figures for more details:

• RFP_OFP_CFP_whiteBG2.jpg bottom of plate on a white background and image processed to reverse the writing
• RFP_OFP_CFP_UVoff2.jpg bottom of plate on UV light (while off) and image processed to reverse the writing
• RFP_OFP_CFP_UV2.jpg bottom of plate on UV light (while on) and image processed to reverse the writing

Here's a chart to summarize how well the protein was produced.

	CFP	OFP	RFP
1	good	bad	good
2	good	good	good

The RFPs are obviously red, even under white light.  The CFPs have the strongest fluorescence under UV light.

We should probably perform some tests to make sure that we are getting the wavelengths that we are expecting.

There seems to be some stray colonies.  This may be from when I was pipetting.

I took the plate out of the incubator at around 9PM on 10/3/14 (about 21 hours in the incubator).

We should NOT use "Orange 1".  If we have time, we could investigate what went wrong.

We should use the rest.

The plates are in the refrigerator (the one furthest back in the lab) at the location indicated by the file plate location.jpg.

Transform 5 ul of each ligation product into commercial chemically competent cells. Incubate at 37C overnight.

 

Look great! Clearly some colonies are fluorescent and some are not (see image)

 

 

Next step: purify the proteins from the cells to ensure they are fluorescent at the correct wavelengths!

add 5ul of digested orange red and cyan fluorescent protein, 7ul of digested puc18, 4 ul of ligase buffer, 1ul ligase, 3 ul of water. leave for the night at room temperature.

We are going to treat the backbone with Antarctic Phoshatase, the protocol in this case is:

1. to the vector tube (which contains 20ul digested pUC18), add 5ul of 10x Antarctic Phosphatase buffer, 25ul of water, and 1 ul of Antarctic Phosphatase. We do this to prevent the vector from closing up again without any insert.

2. Incubate the tube at 37C. After place at 82C for 20 minutes.

 

We assume that the DNA is fully dephosphorylated and that the phosphatase enzyme is denatured.

We need to verify that the fluorescent proteins are fluorescent in the proper color. To do this, we have to first clone the gene (which already has an RBS) into a plasmid with a promoter. This will allow us to express the genes in E. coli for further verification.

Set up digests:

For each fluorescent gene plasmid (OFP, CFP, and RFP. No GFP bc digest looked funny on 9.12:

Digest the plasmid with EcoRI and PstI.

Digest pUC18 with EcoRI and PstI as well. pUC18 is a plasmid with a T7 promoter

Incubate plasmids for 60 min at 37C and then HEAT INACTIVATE enzymes for 20 min at 82C (This last step is very important because we want to use the products of the digests without further purifying them, and we don't want the restriction enzymes to be active when we are putting together the ligations!!!)

Plasmids are digested and ready to use!

Making sure the inserts are here from the colony PCR results - are the GFP and RFP inserts present?

 

Digest each plasmid with EcoRI/PstI. Incubate 15' at 37C.

 

Run entire digest on 0.8% gel.

These were slightly confounding. What I *think* happened is that the buffer wasn't optimal for one of the enzymes, so the digest wasn't run to completion. Why do I think this?

The RFP plasmid should cut to 2.1 kb (backbone) + 0.8 kb (insert). In plasmids 3 and 9, this is the case. 

In the rest of the RFP plasmids, the linearized plasmids appear to be nearly 3 kb. This makes sense if the insert is there but one of the enzymes didn't cut. If the insert weren't present, the backbone should be the 2.1 kb only. Thus, I think they *all* actually have the insert, but we will work with the two confirmed plasmids.

Move forward with RFP 3 and 9.

Followed NEB recommended protocol to transform 5 ul of ligation products into E. coli. We changed the protocol by adding only 40 ul of competent cells (rather than 50 ul as recommended). We also changed the protocol because we wanted to plate all the cells onto the LB+Chlor plates. To concentrate cells, we centrifuged 3 min at 4000 rpm, removed 900 ul, resuspended the cell pellet in the remaining 100 ul, and spread this on an LB+Chlor plate using glass beads.

 

STEPS

1. Thawed a tube of NEB 5-alpha Competent E. coli cells on ice until the last ice crystals disappeared. Mixed gently and pipetted 50uL of cells into a transformation tube on ice.
2. Added 5uL containing 1pg - 100ng of plasmid DNA to the cell mixture. Flicked the tube several times to mix the cells and DNA.
3. Placed the mixture on ice for 30 minutes.
4. Heat shock the mixture for exactly 30 seconds at 42 degrees Celisus.
5. Placed back on ice for 5 minutes.
6. Pipetted 950uL of room temperature SOC with mostly LB into the mixture.
7. Place mixture at 37 degrees Celisus for an hour. Shaked vigorously.
8. Retrieved plates that were warmed at 37 degrees Celisus.
9. Flicked the tube to mix the cells thoroughly. Didn't do any serial dilutions.
10. Spin the cells for 3 minutes at 4,000rpm in the centerfudge. Removed 900mL and then placed the last 100mL onto the plates, using mixing beads, after mixing the cells with the liquid by pipetting up and down. Then we incubated at 30 degrees Celisus for 24hrs.

There was colonies on the plates!

The next step is to check to see if the transformation was done correctly and if E. coli took up the RFP and GFP plasmid.

Digestion of RFP and GFP amplicons using EcoRI and PstI Digest parameters: total volume 20ul H2O: 6ul Buffer(10x): 2ul DNA: 10ul EcoRI: 1ul PstI: 1 ul Made a 1% gel Agarose: 0.5 g TBE: 50ml etBr: 5ul

Miniprep on 8.29.14 of colonies 1-gfp-p, 2-gfp-p, 3-rfp-p, 8-rfp-p, 9-rfp-p, 11-rfp-un, 12-rfp-un, and 16-rfp-un

We have colonies growing from the transformations and want to check if they have the correct plasmid insert. We'll run a colony PCR on these before deciding which colonies to grow and make mini preps from.

 

Ligation	No. Colonies	Notes	Sample Name
BB + GFP (unpur)	0
BB + GFP (pur)	2		1-2
BB + RFP (pur)	7		3-10
BB + RFP (unpur)	7		11-17
BB alone	15	why so many???	18

 

A piece of each of the colonies included in the samples (so not all the BB alone) was inoculated into 20 ul H2O. This was incubated at 95C for 10 min, after which cell debris was removed by a quick centrifugation. 2 ul of the supernatant was used in each of the PCRs.

 

 

PCR set up started by making a cocktail:

Component	ul/rxn	x 18
DEPC-treated water	20	360
Primer 1 = VF2	1.5	27
Primer 2 = VR	1.5	27
Template	2	-
Total	25 (CT = 23)	414

From the CT, 23 ul was aliquoted into each of 18 PCR tubes. 2 ul of the template was added. PCRs were run in the machine with the following protocol:

94C/2min - [94C/30sec  - 55C/30sec - 72C/1min]x25cycles 

Dye was added and the PCR run on a 0.8% agarose gel.

Success!!! From the chart above:

Ligation	No. Colonies	Sample Name	Successful PCRs
BB + GFP (pur)	2	1-2	1,2
BB + RFP (pur)	7	3-10	3,8,9
BB + RFP (unpur	7	11-17	11,12,16

 

We have multiple constructs for both RFP and GFP that may have the correct sequence! 

 

NEXT STEPS:

1. Inoculate the correct colonies into LB+Chlor

2. Miniprep cultures

3. Digest to verify insert size is correct

4. Send for sequence analysis!

 

While waiting for result

transformation: GFP on competent E.coli

Incubate 1 ul plasmid with 50 ul competent E. coli for 30 min on ice 

heatshock 42 C for 45 sec

Incubate on ice 5 min 

Can't fine SOC so we use LB

rotate for 60 mins at 37C

plated on LB+Amp and incubated at 37C

8.21.14: I didn't have time to check the plate yesterday, but two days' growth still looks great - crowded, but still individual, colonies all over the plate. Great!

We want to take Tuesday's PCR products and put them into the pSB1C3 backbone. 

Digest PCRs:

10 ul PCR product

2 ul cutsmart buffer (10x stock)

1 ul PstI

1 ul EcoRI

20 ul total --> incubate for 30 min at 37C

 

PCR purify digest product (only 14 ul - save 6 ul):

Follow kit protocol. Elute in elution buffer.

Worried that the washed columns won't bind DNA, we are going to use some of the set-aside (unpurified) digest product for a backup ligation. We'll run a gel of our purification, but we are going to set up a ligation beforehand, so we won't have even rough estimates of DNA concentrations.

Set up ligations:

Component	Using purified digest product	Using unpurified digest product	BB alone
dH2O	x	11	14
Insert (RFP or GFP)	14	3	x
1:10 BB	3	3	3
T4 buffer (10	1	1	1

The above were incubated 30 min at RT then stored at -20C.

 

We ran a 1% gel of the digest before and after purification. We had a decent yield, maybe 40% of our initial digest product in the purified lanes. 

Next step: Transform the ligated plasmids into E. coli!

RFP prep from colony PCR:

20 µl H2O

inoculate with smidge of bacterial colony

heated 95 C  for 10 mins

spin to remove debris

 

 

PCR; 30 cycles both RFP and GFP

with compositions: 

2 µl template (colony prep or purified plasmid for GFP)

21 µl H2O

1 µl general forward GFP Primer

1 µl specific reversed RFP Primer

 

 

run on 1% agarose gel:

results: good amount of PCR products for both RFP and GFP

RFP #1  

RFP #2 

GFP  

Bands look great! The colony PCR protocol is a real winner!

Next step: clone into pSB1C3 background

RFP prep:

20 µl H2O

heated 95 C  for 10 mins

spin to remove debris

 

PCR: 30 cycles both RFP and GFP

with compositions: 

2 µl template

21 µl H2O

1 µl general forward GFP Primer

1 µl specific reversed RFP Primer

 

run on 1% agarose gel:

results: good amount of PCR products for both RFP and GFP

RFP #1  

RFP #2 

GFP  

 

While waiting for result

transformation: GFP on competent E.coli

Incubate 1 ul plasmid with 50 ul competent E. coli for 30 min on ice 

heatshock 42 C for 45 sec

Incubate on ice 5 min 

Can't fine SOC so we use LB

rotate for 60 mins

plated on LB+Amp and incubated at 37C

The cultures inoculated yesterday are going to be mini prepped by Jonathan. He's also agreed to digest and run a gel of them, to indicate whether the constructs are correct or not.

A. Minipreps were done according to LifeTechnology's protocol. Plasmid products were eluted in 50 ul TE and were used in the digest reactions.

B. Digests were done by first making the following cocktail:

Component	ul/rxn	x 22 rxns
DEPC-treated water	13	286
Cutsmart buffer	2	44
EcoRI	1	22
PstI	1	22

 

From the cocktail, 17 ul was added to 3 ul of mini prep plasmid. This was incubated 15 min at 37C.

 

C. Running the gel. DNA loading dye was added to the digest and the entire digest was run on a 0.8% agarose gel.

More beautiful gels! With more promising results!

The digests were designed to release the fluorescent protein genes so there should be a ~700-800 bp band in addition to a larger band (representing the backbone). 

From A (OFP): Both A1 and A2 are promising and should be sent for sequencing!

From B (YFP): No promising results. If I recall correctly, B is the PCR product that was cut by either EcoRI or PstI (I've lost that gel image!). So it makes sense that the mini preps wouldn't' contain this insert.

From C (RFP): No promising results.

From D (CFP): Lots of promising plasmids! D3, D4, and D6-D9 all have bands of the expected size! I don't know that we need to sequence all of these, but a few should be selected at random and sent for sequencing. 

Both OFP and CFP look to have been cloned into the backbone! Next steps: sequence and submit!

 

We should also start designing a promoter/RBS to add to these plasmids so we can check them for functionality! I'm thinking we could maybe 1. anneal primers that include these sequences and have overlaps with the target plasmid and 2. cut the plasmids at the SpeI site and 3. use Gibson assembly to generate the plasmids!

In his email on Monday, Eric said he saw a few colonies on the ligation transformation plates (the ones Marty plated on 8.10.14):

Name	Construct	# Colonies
BB alone	BB alone
BB + A	OFP	1
BB + B	YFP	2
BB + C	RFP	4
BB + D	CFP	2

Even though there are more colonies on BB alone than BB+insert, I didn't want to throw away our potential constructs. So we will screen these while simultaneously setting up ligations for the next round of transformations. We will screen in two ways: (1) Colony PCR, which will give us results tonight IF IT WORKS, but I've never done colony PCR before and we don't have any positive control primers (meaning that if we see nothing on the gel, we won't know *why* we see nothing on the gel) and (2) traditional mini preps, from cultures we will inoculate tonight into 3 ml LB+Chlor.

From the internet, I gathered several protocols and made an amalgam of them, hoping that this will work to generate template DNA plasmid:

1. Add 20 ul DEPC-treated water to a PCR tube (one per colony to be tested).
2. Using a sterile pipette tip, take *just a smidge* of the edge of the colony. This is the potential biggest source of error, as too many cells or too old of cells will not work. These colonies are old, but if we use the edge of the colony, we can scrape just a bit of the fastest-growing cells (found on the edge) to inoculate into the water.
3. Incubate at 95C for 10 min in the thermocycler.
4. Spin down cell debris on tabletop PCR tube centrifuge (~20 sec). Use 2 ul of supernatant as template DNA in PCR reaction below.

Next, we set up a cocktail for our PCR reactions:

Component	ul/rxn	x 16 rxns
DEPC-treated H2O	20	320
Primer 1: VF2	1.5	24
Primer 2: VR	1.4	24

From this cocktail, we added 23 ul to PCR tubes (containing desiccated PCR  reagents) and 2 ul DNA template, for a total of 25 ul. We ran this in the thermocycler with the following program:

95C/4m - [(95C/30s - 55C/30s - 74C/1m)x30 cycles] - 74C/4 min - 4C/infinity

We added DNA dye directly to the PCR reactions and ran the entire reaction out on a 0.8% agarose gel.

The gel is beautiful! The colony PCR worked!! We can use this screen to decide which cultures need to be mini prepped tomorrow.

 

Here's the bad news: the cultures and PCR reaction numbers are not the same. I can't decipher one from another. I have an *idea* of which is which, but no certainty. This should be avoided in the future with better communication among all team members (I'm not pointing fingers; I just hate that now we have to mini prep all the inoculated cultures. Why for do the short way (colony PCR) if we have to do the long way (mini prep) for all the samples?)

Lanes 1, 3, and 5-15 look very similar in size to one another. These are likely due to backbone ligation with itself in some fashion.

Lanes 2 and 4 look like they have a small insert. This may or may not be the size of one of our fluorescent protein genes. We expect the inserts to be ~700-800 bp *larger* than backbone alone, but since these are consistent in size for the same construct (I think), it's possible they are positive hits.

Lane 16 is the most promising hit. This band shows a shift of about 700 bp above the rest of the bands. This is most likely the A construct, which is OFP. Hopefully mini preps tomorrow confirm this!

We need more GFP because round of ligation did not work, and we have digested GFP. 

We can't find the GFP plasmid to PCR amplify it, so we are amplifying the previously PCRed Digested and Undigested pieces of GFP (instead of the plasmid) hoping that one of them will yield product. 

Two PCR tubes are in the PCR machine. Tube 1D is the digested GFP (left side of machine). Tube 2 is the undigested GFP (right side of machine).

 

We transformed the following ligation products:

• Tube A = OFP
• Tube B = YFP
• Tube C = RFP
• Tube D = CFP (Only had 4 vials of competent cells so did not transform CFP)
• Tube BB = Backbone
• Tube pUC19 = pUC19 Positive Control

1. Added the following to each tube:

Tubes A-D (20 µl total)

• 14 µl PCR product
• 3 µl 1:10 bb
• 2 µl buffer
• 1 µl ligase

Tube BB control (20 µl total)

• 14 µl H2O
• 3 µl 1:10 bb
• 2 µl buffer
• 1 µl ligase

2. Waited 45 mins

3. Pipetted 5 µl of each tube into new tubes labeled A-D and BB.

4. Pipetted 1 µl of pUC19 to pUC19 labeled tube.

5. Add 50 µl of DH5-Alpha to tubes A-D. 

6. Add 1 µl of control DNA to pUC19

Followed NEB 5-alpha Competent E.coli (high efficiency) Transformation Protocol 
(SEE ATTACHED AND BELOW)

1. For C2987H: Thaw a tube of NEB 5-alpha Competent E. coli cells on ice for 10 minutes. 
   For C2987I: Thaw a tube of NEB 5-alpha Competent E. coli cells on ice until the last ice crystals disappear. Mix gently and carefully pipette 50 µl of cells into a transformation tube on ice.
2. Add 1-5 µl containing 1 pg-100 ng of plasmid DNA to the cell mixture. Carefully flick the tube 4-5 times to mix cells and DNA. Do not vortex. (WE ADDED 5 µl)
3. Place the mixture on ice for 30 minutes. Do not mix.
4. Heat shock at exactly 42°C for exactly 30 seconds. Do not mix.
5. Place on ice for 5 minutes. Do not mix.
6. Pipette 950 µl of room temperature SOC into the mixture.
7. Place at 37°C for 60 minutes. Shake vigorously (250 rpm) or rotate. We used rotator inside incubator.
8. Warm selection plates to 37°C.
9. Mix the cells thoroughly by flicking the tube and inverting, then WE DID NOT perform several 10-fold serial dilutions in SOC.
10. Spread 50-100 µl of each dilution onto a selection plate and incubate overnight at 37°C. Alternatively, incubate at 30°C for 24-36 hours or 25°C for 48 hours.

We plated 200 µl from each tube into each plate. Julie will update tomorrow with results.

 

We followed the protocol for Invitrogen Life Technologies One Shot® TOP10 Competent Cells (see attached pdf downloaded on 7/31/14 from invitrogen and excerpt below):

Transform chemically competent cells: Chemical transformation procedure
1. Centrifuge the vial(s) containing the ligation reaction(s) briefly and place on ice.
2. Thaw, on ice, one 50 μL vial of One Shot® cells for each ligation/transformation.
3. Pipet 1–5 μL of each ligation reaction directly into the vial of competent cells and mix by tapping gently. Do not mix by pipetting up and down. The remaining ligation mixture(s) can be stored at −20°C.
4. Incubate the vial(s) on ice for 30 minutes.
5. Incubate for exactly 30 seconds in the 42°C water bath. Do not mix or shake.
6. Remove vial(s) from the 42°C bath and place them on ice.
7. Add 250 μL of pre-warmed S.O.C medium to each vial. S.O.C is a rich medium; sterile technique must be practiced to avoid contamination.
8. Place the vial(s) in a microcentrifuge rack on its side and secure with tape to avoid loss of the vial(s). Shake the vial(s) at 37°C for exactly 1 hour at 225 rpm in a shaking incubator.
9. Spread 20–200 μL from each transformation vial on separate, labeled LB agar plates. The remaining transformation mix may be stored at 4°C and plated out the next day, if desired.
10. Invert the plate(s) and incubate at 37°C overnight.
11. Select colonies and analyze by plasmid isolation, PCR, or sequencing. 

Key Changes we made:

We transformed the following ligation products:

• Tube A = OFP
• Tube B = YFP
• Tube C = RFP
• Tube BB = Backbone
• Did not transform Tube D = CFP (Only had 4 vials of competent cells so did not transform CFP)

For step 3, we pipetted 5 μL of each ligation reaction directly into the vial of competent cells.

For step 7, we did not have SOC media so we substituted LB for SOC media. 

For step 9 we added the following procedure: 

• Spin down 3 mins at 4,000 RPM
• Aspirate and discard 150 microliters
• Resuspend in remaining ~100 microliters
• Plate to LB+Chl

 

 

 

 

No colonies on any plates. Our hypothesis for the poor result is that we used a 1:100 dilution of backbone. It might also be that the cells we were using we not competent.

PCR Purification:

Key for the contents of each tube:

• Tube A = OFP
• Tube B = YFP
• Tube C = RFP
• Tube D = CFP
• Tube BB = Backbone

We used Life Technologies Purelink Invitrogen PCR purification protocol. (See below and attached protocols in pdf format downloaded from Life technologies website on 7/31/14).

PureLink® PCR Purification Kit
Catalog numbers K3100-01 and K3100-02
Publication Part Number 7015021 MAN0004375 Revision Date 14 September 2011

Purifying PCR Products
Procedure for Purifying PCR Products
1. Combine. Add 4 volumes of Binding Buffer B2 or B3 with isopropanol 
(see preceding table) to 1 volume of a PCR sample (50–100 μL). Mix well. 
2. Load. Pipet the sample into a PureLink® Spin Column in a Collection Tube. 
Centrifuge the column at >10,000 × g for 1 minute. Discard the flow-through.
3. Wash. Re-insert the column into the Collection Tube and add 650 µL Wash 
Buffer (W1) with ethanol. Centrifuge the column at >10,000 × g for 1 minute. 
Discard the flow-through and place the column in the same Collection Tube. 
Centrifuge the column at maximum speed for 2–3 minutes.
4. Elute. Place the column into a clean 1.7-mL Elution Tube (supplied with 
the kit). Add 50 µL Elution Buffer to the center of the column. Incubate 
the column at room temperature for 1 minute. Centrifuge the column at 
maximum speed for 2 minutes. The elution tube contains the purified PCR 
product. Store the purified DNA at 4°C for immediate use or at −20°C for 
long-term storage.

We made the following changes to the above protocol: 

1. We started with 20 µL of digest.
2. We used Elution buffer only on tube A.
3. We used deionized Water as buffer for tube B, C, D. The reason for this change is that we felt there might be a conflict between elution buffer and ligation buffer used during ligation process. (For more details ask Julie)

 

Ligation:

14 µL of digested insert (from PCR purification) (Old inserts PCR purified were digested with ecoR1/A+I) (As mentioned above: EB used for A tube. dH2O used for B-D)

3 µL BB (1:100 dilution in H20) (BB=pSBIC3 Ecoli/Ps+I digested on 7-10-14 + PCR purified)

2 µL buffer

1 µL Ligase

------

20 µL total

 

 

Background:

It is possible, albeit unlikely, that the plasmid ends could bind to each other without our gene of interest.  This plasmid still has the same resistance, so the only way to tell that the colonies are different is that they do not express the gene of interest.  With a gene of interest like GFP, it's easy to distinguish between the erroneously formed plasmids and the properly formed plasmids with GFP.  However, this is not always the case.  So, we prepare a control case to be sure that we can count how many erroneous, plasmid-only colonies form.

Ideally, this is the same as the ligation protocol ( https://www.synbiota.com/projects/536/workspace_pages/13142 ;).  

 

 

Background:

vector = plasmid

insert = gene of interest = GFP in this case

Synbiota does something weird to decimal points, so some numbers are represented with spaces and decimal points.

This is what we actually did:

Calculate the appropriate molar ratios (see "Calculating Molar Ratios" https://www.synbiota.com/projects/536/workspace_pages/13144 ).

We decided to use an even higher ratio than 3:1 of gene of interest to plasmid.  We used 6 uL of the gene of interest and 4 uL of the plasmid to help increase the probability of combining the GFPs to the plasmids

We need to sum to 20 uL of solution total.

  9 uL of deionized H20

  4 uL of digested plasmid

  6 uL of PCR purified GFP

  2 uL of 2x ligation buffer

  1 uL of ligase

~20 uL total (actually 21 uL)

This is from the NEB website https://www.neb.com/protocols/1/01/01/dna-ligation-with-t4-dna-ligase-m0202 :

COMPONENT	20 μl REACTION
10X T4 DNA Ligase Buffer*	2 μl
Vector DNA (4 kb)	50 ng (0 . 020 pmol)
Insert DNA (1 kb)	37.5 ng (0 . 060 pmol)
Nuclease-free water	to 20 μl
T4 DNA Ligase	1 μl

* The T4 DNA Ligase Buffer should be thawed and resuspended at room temperature.

This is what we actually did:

1. Combine the materials in a PCR tube in THIS ORDER:
   1. H2O
   2. buffer 
   3. DNA (plasmid and GFP, the order does not matter)
   4. enzyme (ligase)
2. Mix with pipettor.
3. Incubate at 16C in the PCR machine.  Actually was at 16C for 24 minutes then room temperature for about 23.5 hours.
4. Put on ice in the freezer.

This is from the NEB website https://www.neb.com/protocols/1/01/01/dna-ligation-with-t4-dna-ligase-m0202 :

1. Set up the following reaction in a microcentrifuge tube on ice.
   (T4 DNA Ligase should be added last. Note that the table shows a ligation using a molar ratio of 1:3 vector to insert for the indicated DNA sizes.) Use NEBioCalculator to calculate molar ratios.
2. Gently mix the reaction by pipetting up and down and microfuge briefly.
3. For cohesive (sticky) ends, incubate at 16°C overnight or room temperature for 10 minutes.
4. For blunt ends or single base overhangs, incubate at 16°C overnight or room temperature for 2 hours(alternatively, high concentration T4 DNA Ligase can be used in a 10 minute ligation).
5. Chill on ice and transform 1-5 μl of the reaction into 50 μl competent cells.

 

Background info:

• The plasmid is about 2000 base pairs (bp).  
• Our PCRed GFP (gene of interest) is about 750 bp.
• Assume that 1 uL of this PCR purified GFP = 50 ng (info from Ellen).

Calculations:

We have 6 uL of our gene of interest in a solution totaling 20 uL.

So we have:

\(\displaystyle \frac{6\mu L }{1}PCR \; purified \; GFP * \frac{50 ng}{1 \mu L} * \frac{1}{20\mu L} = \frac{300 ng}{20 \mu L} PCR \; purified \: GFP = 15\frac{ng}{\mu L}PCR \; purified \; GFP\)

 

The iGEM kit provides a linearized plasmid backbone in a solution with a concentration of 25 ng/uL.  

In our digestion step, we used 4 uL of the linearized plasmid backbone and 4 uL of the enzyme master mix. 

So, we have:

\(\displaystyle \frac{4\mu L }{1}plasmid * \frac{25 ng}{1 \mu L} * \frac{1}{(4+4)\mu L} = \frac{100 ng}{8 \mu L} plasmid= 12.5\frac{ng}{\mu L}plasmid\)

We need to have a GFP:plasmid ratio of at least 3:1 to make sure that we have enough pieces of the gene of interest to successfully connect to the plasmid backbone.

Now, we'll use the NEBioCalculator  to get the number of moles for each (see the pictures).

So, we have:

\(\displaystyle \frac{32 . 36 \frac{fmol}{\mu L} \; GFP}{9 . 632 \frac{fmol}{\mu L} \; plasmid} = 3 . 36 \; GFP : 1 \; plasmid \gt 3 \; GFP : 1 \; plasmid\)

Therefore, we can use a ratio of  1 uL of the PCR purified GFP solution to 1 uL of the digested plasmid solution.

 

DNA from mini prep

10x buffer designed to work with enzyme

enzyme

dH2O

thermal cycler

gel box

agarose

1x TAE

ethidium bromide

 

for a 20ul reaction: 

 for one reaction tube:

reagent	ul
dH2O	12
buffer (10x)	2
enzyme	1
DNA	5

total volume

20

1. In a 25ul PCR reaction tube, add 12 ul of dH2O, 5ul of 10x buffer (buffer must be compatible with enzyme being used), add 5 ul of DNA product, and lastly add 1 ul of enzyme.

* if making a number of tubes, create a cocktail with enough reagents to aliquot into all tubes used. combine dH2O, buffer, and enzyme into a 2ml tube. aliquot 15ul of cocktail into 25ul PCR reaction tubes. Add DNA directly to PCR tubes. 

2. cover tubes and place in thermal cycler and run restriction digest protocol

3. after the run is completed, follow prcedure 3: running a gel in order to visualize the results of your restriction digest

 

1.50ul vial of top 10 cells

2. DNA of desired plasmid

3. ice block

4. pipette

5. water bath (set to 42oC)

6. Vial of SoC

7. Shaker

8. plate with LB + limiting factor (for this it is Chlore)

9. Incubation chamber set to 37oC

1. thaw on ice one 50ul vial of one shot top 10 cells for each reaction you will be performing

2. add 2ul of plasmid DNA to the vial. mix by tapping gently. do not pipette up and down

3. incubate on ice for 30 minutes

4. heat shock the tubes for 30 seconds at 42oC

5. incubate cells on ice immediately after heat shocking for 1-5 minutes

6. off ice, add 250ul of Soc to the reaction tube. shake for 1 hr at 37oC

7. spread 10-50 ul of the reaction to pre-set LB+ chlore plates

8. incubate overnight at 37oC

• 9.5 µL of deonized water
• 6 µL of GFP DNA
• 1 µL of EcoRI (restriction enzyme)
• 1.5 µL if PstI (restriction enzyme)
• 2 µL of buffer (must match restriction enzymes used – check with lab director)

IMPORTANT: Add materials in this order:             

• Water
• Buffer
• DNA
• Restriction Enzymes

Flick to mix.

Heat in PCR machine at 37 degrees C for 1 hour

Ice mixture

*BREAKPOINT*

Run a Gel

Now that we have a purified PCR produce we would like to confirm that we actually have some DNA that appears to be correct.  To do this, we will run a gel.  GFP contains approximately 783 base pairs so we will run 100 bp ladder to compare the PCR product to.

The first step is to make a gel of the right concentration.  Gels concentrations run 0.7% - 2%.

For 500 bp sequences, concentration of > 1% is good.  For 783 bp, a 1% gel should work.

Make the gel:

Materials:

                0.5 g of agarose

                50 mL of TAE

                5 µL of Ethidium Bromide (for small gel)

                Small Electrophoresis box

Step 1:  Measure .5 g of agarose.  Use folding paper (fold in quarters first).  Also be sure to take scale after putting on folding paper.

Step 2:  In 250 mL Erlenmeyer Flask, mix agarose and 50 mL of TAE.  Swirl flask to dissolve most of the agarose.  Liquid will be foggy.  Cover flask with Saran Wrap and microwave for 2 minutes at high.  Liquid will be clear.

Step 3:  Add 5 µL of Ethidium Bromide.  Swirl.

Step 4. Set up electrophoresis box to make gel.  Make sure tray is oriented so that liquid gel will not leak out.  Insert well mold.  Pour in enough gel mix to fill tray.  Let sit for about 15 minutes (will turn foggy when ready) to allow gel to set.  Rinse flash in tap water.

Step 5:  Add a splash of TAE to lubricate gel.  Wiggle out well mold.  Gently take out tray and rotate 90 degrees.  Make sure that wells are on NEGATIVE side (black lead) or sample will run the wrong direction (DNA is negative and will be drawn to the positive side).

Step 6:  Fill gel box with TAE 1% solution.  It should just go over the top of the gel.

Step 7:  Get DNA sample ready to run gel.  Place a small piece of parafilm over a test tube tray and make a small indentation (area will be used to mix DNA and loading dye).  Mix 2 µL (we used 9 in error) of DNA sample with 1 µL of loading dye in parafilm indentation.  Using pipette, place full mixture into one of the gel wells.  In a neighboring well, place in 3 µL of 100 base pair ladder.

Step 8:  Run the gel by plugging leads into the transformer.  If power is on, you will see small bubbles coming off of wires.  Let the gel run until the dye has gone about 1/3 the way across the gel.

Step 9:  To see the actual DNA, take the gel tray and view it under ultraviolet light.  If everything worked you will see one band in the DNA column and multiple bands in the ladder column.  See where the DNA band lines up against the ladder to estimate the number of base pairs and to see if they correspond to the expected amount.  Take a picture of the gel.

PCR Purification

In this next step we need to remove the polymerase from the test tube.  A later step in this process will be to convert the blunt ends of the GFP DNA into sticky ends.  If the polymerase is not removed then it will be impossible to create the sticky ends since as the sticky ends are created, the polymerase will convert them back to blunt ends.  The purification process is done with a kit.  In our lab we used the PureLink Quick Gel Extraction and PCR Purification Combo made by Invitrogen (Catalog number: K2200-01)

Step 1:  Combine. Add 4 volumes of Binding Buffer (B2) with isopropanol to 1 volume of PCR sample (in our case, 88 µL of B2).  Mix well.

Step 2:  Load.  Pipet the sample into a PureLink Clean-up Spin Column in a Wash Tube.  Centrifuge the column at >10,000 x g for 1 minute.  Discard the flow through.

Step 3:  Wash. Re-insert the column into the Wash Tuben and add 650 µL Wash Buffer (W1) with ethanol.  Centrifuge the column at >10,000 x g for 1 minute.

Step 4:  Remove ethanol.  Discard the flow-through and place the column in the same Wash Tube.  Centrifuge the column at maximum speed for 2-3 minutes.

Step 5:  Elute.  Place the column into a clean 1.7 mL Elution Tube.  Add 50 µL Elution Buffer (E1) to the column. Incubate the column at room temperature for 1 minute.  Centrifuge the column at maximum for 1 minute.

The elution tube contains the purified PCR product.  Make sure to label the tube.

*BREAKPOINT*

• Forward & Reverse Primers

• PCR bead in tube (bead contains polymerase and nucleotides)

• 19 µL of deionized water (we used 16 µL by mistake and it still worked)

• 1 µL of template plasmid

• 2.5 µL of 10x forward primer solution

• 2.5 µL of 10x reverse primer solution

PCR Primer DNA

Forward and reverse primer DNA are shipped in separate storage tubes.  The first step is to make a 100X solution of each in the storage tubes by adding deionized water. 

Add the appropriate amount of water and set aside.

Make 10x solution of the primer DNAs

In separate small Eppendorf tubes, mix 10 µL of the primers with 90 µL of deionized water.  Close top and vortex each tube.  Make sure to label the tubes.  Set aside.

PCR Reaction

The objective in this step is to clone the DNA in the template plasmid (GFP in our case).

Step 1: Add deionized water to PCR tube.  Flick tube to dissolve PCR bead. (DO NOT VORTEX – amount is too small.)

Step 2:  Add template plasmid, forward primer, and reverse primer.  Flick tube to mix (DO NOT VORTEX)

Step 3:  Place in PCR machine and run through cycle (takes about 90 minutes)

Step 4:  Remove & label tube

If all went well, you should have a tube of lots of the template DNA with blunt ends.

*BREAKPOINT* (A breakpoint is a place where you can stop working.  Please make sure all of your test tubes are labeled and placed in the freezer in the iGEM box.)

I inoculated 12.5 ul of the overnight culture into 25 ml YPD at 11pm Tuesday night. On Wednesday night, the culture was already dense. I didn't measure OD however. 

The protocol is fairly straightforward. During the trial run, I encountered a few hurdles (e.g., fuse of centrifuge went out with samples still inside; had to buy a new fuse; samples were left pelleted in the centrifuge for ~30 extra minutes) but this is overall a permissive protocol with a lot of wiggle room for timing changes. 

Prepare ahead of time: 

TELiOAc: 10 mM TE pH 8.0, 0.5 mM EDTA, 100 mM LiAc

PLATE: 40% final concentration PEG 3350, 10 mM TE pH 8.0, 0.5 mM EDTA, 100 mM LiAc

Steps:

1. Collect an exponentially growing culture of S. cerevisiae by centrifugation (I used the W303A strain). Yeast cells are pretty big and don't need to be spun as quickly as bacteria to sediment; I used 5 min at 4000 rpm in the IEC tabletop centrifuge.

2. Wash cell pellet by resuspending in TELiOAc. Use 5 ml TELiOAc for every 50 ml culture grown (I grew 25 ml culture, so 2.5 ml TELiOAc). Pellet cells again.

3. Resuspend cells in TELiOAc. Use 0.5 ml TELiOAc for every 50 ml culture grown (I used 0.25 ml, the same as saying 250 ul).

4. Prepare 1.5 ml epitomes with DNA. In this case, I have three samples. Each tube gets 5 ul carrier DNA (we're using salmon sperm DNA). Two tubes get 5 ul EcoRI-digested and gel purified pACT-AD. One of these tubes gets 20 ul HindIII-digested and gel purified pACT-AD. (See table in results section)

5. Add 100 ul yeast cell suspension. Incubate at room temperature for 30 minutes.

6. Add 0.5 ml (500 ul) PLATE mixture to each sample

7. Heat shock 5 min at 42C.

8. Pellet cells gently by spinning 3 min, 4000 rpm. Discard supernatant. Gently wash off cell pellet with 1 ml YPD *WITHOUT DISRUPTING THE PELLET*. Resuspend pellet in 100 ul YPD and spread over selective plates (sc-leu in this case).

9. Incubate at 30C for two days. Inspect plates for colonies. (NOTE: The thermometer in the nonAC part of Genspace reads 29C -- this is close enough. I suspected Genspace would be great for growing yeast!)

Conditions	carrier DNA alone	c. DNA + backbone	c. DNA + BB + insert
Expected CFU	none	few	hundreds
Observed CFU

I'll leave the plates incubating until Friday or possibly Saturday and fill in the results soon!

I want to test out the yeast strains and shuttle vectors that we hope to use - need to make sure we can get the transformation to work. 

A short bit on the theory behind in vivo recombination:

Homologous recombination works similar to Gibson assembly, insomuch as homologous ends promote recombination. We put linear pieces of DNA into Saccharomyces cerevisiae - this the the same yeast that can make beer and bread (although those require different strains). S. cerevisiae doesn't like short, linear pieces of DNA, but it is really good at turning linear pieces of DNA into circular plasmids. If the ends of the DNA are the same, the yeast DNA repair enzymes will target these to be integrated. 

I have two plasmids: p426gpd (URA marker, 6606 bp) and pACT-AD (LEU marker, 8117 bp). These plasmids allow certain strains of yeast to grow in the absence of uracil or leucine, respectively. (This is a different selection method than antibiotic resistance). I'm going to test the in vivo recombination protocol using the pACT-AD plasmid.

 

6.30.14 

Start cultures of E. coli containing each plasmid

 

7.1.14

Miniprep E. coli to recover plasmids. Make a glycerol stock of each strain (can be found in iGEM 2014 box). Store mini prepped plasmids in iGEM 2014 box too.

Digest the mini preps:

Digest 1: 

12 ul H2O

5 ul pACT-AD

2 ul NEB Buffer 2 (WRONG BUFFER FOR PROMEGA ENZYME)

1 ul HindIII

 

Digest 2: 

11 ul H2O

5 u pACT-AD DNA

2 ul Cutsmart buffer

1 ul EcoRI-HF

1 ul BamHI-HF

 

Digest 3: 

14 ul H2O

2 ul NEBuffer 2

3 ul pACT-AD DNA

1 ul EcoRI

 

Digest 4: 

14 ul H2O

2 ul NEBuffer 2

3 ul p426gpd DNA

1 ul EcoRI

 

Incubate all at 37C for ~1 h.

Run on 0.8% agarose gel.

 

Results: 

Digest 1: Great! HindIII worked even though the enzyme was from promega and I used the NEBuffer 2. Cut out circled 1 kb band

Digest 2: Terrible! Where did the DNA go? Either the BamHI or Cutsmart buffer are no good, methinks

Digest 3: Great! Linearized pACT-AD --> I'm going to use this because Digest 2 didn't work. Cut out to purify

Digest 4: Strange! The pattern is not what I expected. I thought to see a single 6.6 kb band but I see a 6.6 kb and something else - I bet it's an insert in the plasmid. I'm going to cut out the 6.6 kb band to gel purify as well.

DNA is ready to go! I'm transforming the S. cerevisiae tomorrow. Will start a culture tonight.

We want to move some Biobrick parts into E. coli. The parts we've selected are:

Description	Part	Location	Size (bp)	Performed by
Super YFP 2	Bba_K864100	2014 1-17B	723	Ramsez
HSP Promoter	BBa_K873002	2014 1-3H	47	Christal
Apple Fragrance Generator	BBa_K395602	2014 1-4H	1813	Eric
Luciferase	BBa_K325210	2014 1-2B	2623	Anne
Epic luciferase	BBa_K325108	2014 1-2F	2891	Cecil

pSB1C3 2070 bp

We did a quick-and-dirty transformation into Top10 E. coli cells (see protocols for more details).

For Luciferase, Epic luciferase, and Apple Fragrance Generator, we plated both a 1:10 dilution and the rest of the cells on two separate plates (after concentrating the remaining 450 ul). The HSP promoter and Super YFP were only plated in 1:10 dilutions.

Only the SYFP and Apple Fragrance Generator constructs produced colonies. They were inoculated into LB + Chlor (thanks StuyGEM!) for culturing, mini preps, and digestion mapping.

• 50 mM CaCl2
• Eppindorf tubes
• A plate of E. coli
• LB
• Ice
• LB agar plates with selectable drug

1. Add 250 ul of 50 mM CaCl2 to an eppindorf tube.
2. Scrape E. coli from plate and add to CaCl2
3. Incubate 15-30 min on ice
4. Add DNA (in case of biobrick parts, resuspend in 10 ul and add 2 ul to cell suspension)
5. Heat shock 42C for 45 sec
6. Incubate 1-2 min on ice
7. Add 250 ul LB. Incubate 20 min @ 37C
8. Plate on LB + selection drug

Long-awaited and finally here…a bit on primer design and usage. Thought it might be helpful before our meeting on Wednesday!

 

Julie

We might incorporate some sort of visual like this on our wiki (I'm sure the designers can make it look jazzier!)

In our quest to build an IP-free plasmid, we may take shortcuts by using pieces of available plasmids. These papers outline a little bit about the history of these plasmids, which were originally generated using a bacteriophage (a virus that infects bacteria) cloning system.

These papers help to trace back the origins of the pUC18/19 plasmid system. pUC18 and pUC19 are the same plasmid, with one sequence area of the plasmid (the multiple cloning site) reversed between the two. They are commercially available and we have them in Genspace. The origins of these plasmids are quite old, with the literature describing them going back to the 70s and 80s. This likely indicates that intellectual property is probably not a big hurdle if we want to use certain sequences from these plasmids.

 

The pUC plasmids, an M13mp7-derived system for insertion mutagenesis:

- Geneology of some pUC plasmids (not pUC18/19 but similar in certain aspects). (See Fig 1)

 

Cloning in M13 phage or how to use biology at its best:

- A first-hand historical account of the experiments to make the ancestors of the pUC18/19 plasmid systems.

- Contains a useful table that illustrates the history of some of these plasmids/phages.

While a great strength of iGEM is the interdisciplinary nature of the teams that compete, it is still crucial for every member of a team to have a basic grounding in synthetic biology concepts and wet work skills. 

This is a list of biology concepts and skills that everyone on an iGEM team should know. 

Credit for the seed of this list to Julie Wolf (Thanks Julie!)

Synthetic Biology Concepts

1. DNA
   1. double-stranded
   2. bidirectional
   3. complementary
2. Nucleotide (NT)
   1. basic building block
   2. oligomer = several NTs
3. Gene
   1. designated set of NTs
   2. turned into mRNA
   3. product is usually a protein 
4. PCR
   1. generates many copies of DNA in vitro (no cells required)
   2. requires oligos to act as primers for the DNA polymerase enzyme 
5. Restriction Enzymes (playlist)
   1. Discovered in bacteria to fight invading bacteriophage
   2. cleave DNA at specific sites in specific patters
      1. sticky v. nonsticky ends
6. ​Ligase (vid)
   1. Ties together phosphate backbone to covalently bond two DNA pieces
   2. Necessary to make ends tie together
7. Gibson Assembly (vid)
   1. takes advantage of three-enzyme cocktail (an endonuclease, a polymerase, & a ligase)
   2. greatest advantage: we don't depend on restriction sites for where to tie DNA together
8. Yeast homologous recombination mechanism (in vivo) 
   1. better than Gibson Assembly (? - Yuriy)

Lab Skills

I found these playlists to be helpful. I don't know what it will be like to a novice. There is another by bioorad. I wish these were around when learning about restriction enzymes (Yuriy)

I found a class from MIT on youtube that also explains these concepts.  You can access it here: MIT 7.01SC Fundamentals of Biology.  The class is more extensive than just this outline so you can either watch the whole class or just specific lectures (Marty)

Tools of the trade

• micropipettor (video Standards Of Practice)
•  

 

 

Extend an invitation to people who taken classes at Genspace.

Maybe name change.

What's up with the moss

 

A detailed description of the mechanism of action of Colicin V.

The article that describes the details of the colicin V system.