Team:UChicago/Notebook

Contents 1 June 1.1 Week 3 1.1.1 Tuesday, June 18, 2013 1.2 Week 4 1.2.1 Wednesday, June 26, 2013 1.2.2 Friday, June 28, 2013 2 July 2.1 Week 1 2.1.1 Wednesday, July 3, 2013 2.1.2 Thursday, July 4, 2013 2.1.3 Friday, July 5, 2013 2.1.4 Saturday, July 6, 2013 2.2 Week 2 2.2.1 Monday, July 8, 2013 2.2.2 Wednesday, July 10, 2013 2.2.3 Thursday, July 11, 2013 2.3 Week 3 2.4 Week 4 2.4.1 Tuesday, July 23, 2013 2.4.2 Wednesday, July 24, 2013 2.4.3 Friday, July 26, 2013 2.5 Week 5 2.5.1 Monday, July 29, 2013 2.5.2 Tuesday, July 30, 2013 2.5.3 Wednesday, July 31, 2013 3 August 3.1 Week 1 3.1.1 Thursday, August 1, 2013 3.1.2 Friday, August 2, 2013 3.1.3 Saturday, August 3, 2013 3.1.4 Sunday, August 4, 2013 3.2 Week 2 3.3 Week 3 3.3.1 Tuesday, August 20, 2013 3.3.2 Wednesday, August 21, 2013 3.3.3 Thursday, August 22, 2013 3.3.4 Friday, August 23, 2013 3.4 Week 4 3.4.1 Monday, August 26, 2013 3.4.2 Tuesday, August 27, 2013 3.4.3 Wednesday, August 28, 2013 3.4.4 Thursday, August 29, 2013 3.4.5 Friday, August 30, 2013 4 September 4.1 Week 1 4.1.1 Tuesday, September 3, 2013 4.1.2 Wednesday, September 4, 2013 4.1.3 Thursday, September 5, 2013 4.1.4 Friday, September 6, 2013 4.2 Week 2 4.2.1 Monday, September 9, 2013 4.2.2 Tuesday, September 10, 2013 4.2.3 Wednesday, September 11, 2013 4.2.4 Thursday, September 12, 2013 4.2.5 Friday, September 13, 2013 4.2.6 Saturday, September 14, 2013 4.3 Week 3 4.3.1 Monday, September 16, 2013

June

Week 3

Tuesday, June 18, 2013

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First iGEM Meeting of the Summer!

• All student members gathered in the lab.

• We toured the lab, prep room, and incubator room, and we discussed the lab rules.

• We also split into teams to discuss lab logistics: contacting funding sources, ordering reagents/materials, compiling protocols, recording our electronic lab notebook, and making plates/media.

Week 4

Wednesday, June 26, 2013

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Planning Wet Lab Work

• We split into planning sub-teams to plan out different parts of the project with protocols, timelines, and ordering reagents.

• Plasmid Design
• Media and Competent Cells
• E. coli and B. subtilis Transformation
• Keratinase Assay and His tagging
• Running DNA gels, SDS-PAGE gels, and western blots

• We will also be holding weekly meetings with our graduate student advisers and biweekly meetings with our faculty advisers to provide updates.
• Wet lab work begins tomorrow!

Friday, June 28, 2013

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Researching the Sequence of kerA

• Planning subteams continued to order reagents and compile protocols.
• The plasmid design team decided on gibson assembly to produce the kerA biobrick with geneblocks ordered from IDT.
• kerA sequence was obtained from “Nucleotide Sequence and Expression of kerA, the Gene Encoding a Keratinolytic Protease of Bacillus licheniformis PWD-1” (Lin et al. 1995).

• Two putative promoters (red), a possible ribosomal binding site (green), and a transcriptional terminator sequence (light blue, single underline) are indicated. The putative starting residues of the preprotein (pre = blue), proprotein (pro = pink), and mature protein (mature = purple) are indicated.
• EcoRI (orange) and PstI (blue) sites within the gene are indicated and must be removed to produce the kerA biobrick.

July

Week 1

Wednesday, July 3, 2013

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BioBrick Design Presentation and Grad Adviser Meeting

• Designed a biobrick with:

• C terminus His-tagged kerA

• CATCATCATCATCATCAT

• Prefix: gaattcgcggccgcttctagag
• Suffix: tactagtagcggccgctgcag

• Moreover, the bio brick must include

• Eliminated EcoRI and PstI restriction sites from kerA nucleotide seq

• EcoRI (gaattc--814) and PstI (ctgcag--978)
• Conserved aa seq + biobrick prefix + suffix

• Secretion signal peptide
• Inducible promoter

• We also presented a feasible timeline of our project, which included:

• Assembly of biobrick and transformation into E. coli
• Transformation of B. subtilis
• Keratinase isolation and keratinase assays
• kerA mutagenesis for optimization of the keratinase

• We also split up basic lab duties!

• Ivan & Ethan are in charge of the IBC protocol.

• We can’t start wet lab work until our IBC protocol is finished!

• Ethan is in charge of minutes.
• Susan is in charge of ordering invoices.
• Annie is in charge of emailing advisors and members to plan meetings
• Chloe is getting the B. subtilis gene that expressing keratinase.

Thursday, July 4, 2013

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Milk Agar Plates

• The purpose of milk agar plates is often to detect protease activity, including the activity of keratinases.
• A measurement of any baseline protease activity of our parent B. subtilis strain (WB700), which does not endogenously express any keratinases, is desired.
• Excess protease activity of our parent B. subtilis strain may prevent our use of milk agar plates as an assay for keratinase expression of our future transformants with the kerA gene.
• Therefore, the parent B. subitilis strain (WB700) was streaked on the milk agar plates to observe baseline protease activity that could prevent us from detecting keratinase expression after transformation.
• Milk Agar Plates (in LB):

• Total volume of 600 ml (makes 20-25 plates)
• 20g (30*.6) nonfat dry milk powder
• 500mL 2X LB medium
• 250mL 2X agar

• Ideally, there should be a minimal zone of clearing around our B. subtilis colonies.
• Next Steps:

• Observe the results of our milk agar plates with B. subtilis colonies

Friday, July 5, 2013

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Primer Design for kerA biobrick

• Designed primers for directed mutagenesis of our kerA biobrick using [http://www.bioinformatics.org/primerx/ PrimerX]:

1. Primer pair 1 (EcoRI mutation, aat --> aac, asparagine)

   a. Forward: 5' GGTTAAAGTACTGAACTCAAGCGGAAGCGG 3'

   b. Reverse: 5' CCGCTTCCGCTTGAGTTCAGTACTTTAACC 3'

2. Primer pair 2 (Pst1 mutation, gca-->gcg, alanine)

   a. Forward: 5' GTTGTCGTTGTAGCTGCGGCAGGGAACAGCGGATC 3'

   b. Reverse: 5' GATCCGCTGTTCCCTGCCGCAGCTACAACGACAAC 3'

• Used APE for checking codons and mutating restriction sites

Saturday, July 6, 2013

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Primer Design Part II

• Designed primers to isolate non-coding and coding sequence of kerA and add prefix and suffix

• forward primer + xbaI site:

5' TATCTAGAGGTCTATTCATACTTTCGAA 3'

• reverse primer + speI site (reverse complement):

5' ATTGATCATTGGACAACCTTCATCAG 3'

• We still need to design additional primers that will include the secretion signal peptide (in case it needs to be secreted out of B. subtilis) and kerA (From start to stop codon).
• These primers should include igem biobrick prefix and his tag (before stop codon) + suffix.

Week 2

Monday, July 8, 2013

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Milk Agar Plate Results

• The results of our milk agar assay from 7/04/13 of the non-transformed B. subtilis strain WB700:

• The zone of clearing around the B. subtilis colonies indicates that our parent strain breaks down milk. Its prominent endogenous protease activity prevents us from using the milk agar assay as a screen for successfully transformed kerA-expressing B. subtilis.

• Alternative method to screen for transformed B. subtilis needed:

1. The feather assay used in Saha and Dhanasekaran’s 2010 paper, “Isolation and Screening of Keratinolytic Actinobacteria form Keratin Waste Dumped Soil in Tiruchirappalli and Nammakkal, Tamil Nadu, India”
2. Quantitative assay using azokeratin

Our future plans:

• For our promoter for kerA, Karyl, one of our graduate advisors, recommended P43, a constitutive promoter, to transform into B. subtilis.

• We also decided to use pUB110, a high copy number plasmid used in S. aureus and B. subtilis, to introduce kerA into our parent B. subtilis strain.
  • pUB110 will also be made into a biobrick.

• We have to choose a suitable plasmid (either pSB1A3, pSB1C3, pSB1T3, or pSB1K3.m1) for our E. coli transformations.

Wednesday, July 10, 2013

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Grad Advisor Meeting on further developed Keratinase Project

• We presented the primers we designed to our advisers, who suggested we double check them for self dimerization and 3’ overlaps with:

• • http://www.idtdna.com/analyzer/applications/oligoanalyzer/
  • http://bioinfo.ut.ee/primer3/

• We discovered that our forward primer forms a self-dimer and must be redesigned.

• We also need lysozyme to digest the peptidoglycan of B. subtilis in order to miniprep it.

• We decided to use the iGEM plasmid pSB1K3.m1 for our E. coli transformations.

• To screen for keratinase activity, instead of using the feather assay, which takes up to a month for visible results to appear, we decided to use the azokeratin assay as the secondary screening method.

Thursday, July 11, 2013

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Meeting with Purdue iGEM team

Get Excited for the UChicago X Purdue collaboration! Our first meeting with the Purdue team went well:

• The Purdue team will send us a form to provide feedback on their igem biobrick characterization and protocol forms on Monday.

• The biobrick forms are in PDF format.

• Their biobrick characterization contains info like biobrick sequence, plasmid and microorganism in which it’s inserted, ect.

• The protocol form is a checklist of the protocols carried out to characterize the biobrick. It doesn’t contain much info.

• Nora asked them to add a section stating the purpose for using a specific protocol, as in, what specifically about the biobrick are they trying to characterize.

Week 3

Week 4

Tuesday, July 23, 2013

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Today we made LB media with kanamycin and chloramphenicol. There are 7 plates of each kind of antibiotic marker left over; we put them in the 4ºC fridge. We autoclaved all the media that we made today and yesterday.

We transformed the plasmid with promoter Veg (in the pSB1C3 vector) into TOP10 E. coli competent cells. We got the DNA (part BBa_K143012) from well 23B on the distribution kit. The bacteria have been plated onto a chloramphenicol plate. They have been left to incubate at 37ºC in Nora’s lab (GCIS W519).

Future plans:

• Select a few colonies and inoculate overnight in 3 mL cultures (These will eventually be used to make glycerol stocks and for DNA miniprep). Do this later on in the day... ~3 pm onwards.
• Store the plate in 4ºC afterwards in case it is needed in the future.
• Make liquid LB media (no agar!) with 50 µg/mL chloramphenicol.

Wednesday, July 24, 2013

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2 colonies were selected from yesterday's pVeg transformants to be grown in an overnight liquid culture. However, we redid the transformation using a slightly different procedure because there were very few transformants:

1. Let the cells thaw on ice.
2. Add your plasmid to the competent cells.
3. Leave on ice for 10 mins.
4. Place on the 42ºC heat block for 45 s to heat shock.
5. Put on ice for 2 minutes.
6. Add 1 mL SOC media.
7. Incubate in the 37ºC shaker for 1 hour.
8. Centrifuge at 8 rpm for 30 seconds. Decant all but ~100 µL supernatant and resuspend.
9. Plate onto chloramphenicol (or antibiotic of choice) plates and use glass beads to spread out the colonies.

Future plans:

• Make DH5a competent cells; our poor transformation results could be due to incompetent cells.
• Isolate Pveg (for B. subtilis) promoter from transformants grown overnight

• Do DNA miniprep of the transformants
• Cut w/ EcoRI and SpeI
• Run in 0.9% or 1% agarose gel (make gel w/ large wells, load 50 uL of sample)
• If there is a band of right size of Pveg, do a gel extraction
• Put at -20ºC
• If there is no correct band, set overnight cultures of transformants from the transfromation done by Ivan today (7/24/13)

• And for Friday's meeting (7/26/13)

• Isolate Pveg (for B. subtilis) promoter as described above if it was unsuccessful the day before. Use the overnight cultures set up on 7/25/13.
• Check if the transformation w/ the E. coli constitutive promoter was successful. If yes, store plate at 4ºC.

Friday, July 26, 2013

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To do:</i>

• Make overnight cultures of colonies from Annie’s transformation.
• Transform 18O and 18E again, this time into the competent cells made yesterday by Ethan and Annie. Those plasmids should be in the -20ºC.--DO MONDAY, Ethan’s doing with transformation efficiency 7/29
• Check the efficiency of our competent cells by using the transformation efficiency kit from iGEM (check the instructions on the iGEM wiki).--DO MONDAY
• Do a gel purification of the digestion of the PVeg promoter & check the size.

• The digested DNA is sitting in the fridge. Use ALL of it.

Week 5

Monday, July 29, 2013

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<i>To do:

• Set O.N. culture of 18O (2X 3mL)--done by Annie (18O is the E. coli constitutive promoter)
• Check transformation efficiency, 18E of competent cells made by Ethan
• Run gel with Pveg digestion (cut w/ EcoRI and SpeI)
• Redo the digestion with 10uL of DNA miniprep of Pveg w/ EcoRI and SpeI for 2 hr
• Run in 2% agarose

• If there are bands--purify promoter
• If there are no bands, set up O.N. culture of Ivan’s transformation of Pveg

• Set O.N. culture of B. subtilis BD 366 w/ pUB110 (2X 3mL)--done by Nora

Getting done today:

• Re-digest pVeg, digest 5ul of miniprep 1 2hrs at 37 deg C:

• 2 ul 10X buffer 2.1
• 1 ul EcoRI HF
• 1 ul SpeI
• 5 ul mini prep 1
• H2O to 20 ul

• Also set previously done digest by Klevin for 2 hrs at 37 deg C

--Done by Annie and Nora

• Transformed 50 uls of new competent cells and old competent cells with 0.5 ul of 18E resuspension and plated 50 ul of new competent cells as a negative control

--Done by Ethan

Next Steps:

• Run digest of Pveg on 2% agarose gel
• Do miniprep of B. subtilis pUB110
• Do miniprep of 18O transformants

Tuesday, July 30, 2013

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• Ran gel of Pveg miniprep digest (Klevin’s at total 3hrs, and Nora’s/Annie’s at 2hrs digestion) with EcoRI and SpeI

• 10 ul ladder
• 20 ul of samples + 5 ul of Orange G dye

• Final gel: Nora’s and Annie’s (N&A) had faint band at 100 band marker, we assume this is the fragment
• Nora purified gel fragment, with Qiagen kit, eluted in 30ul water in order to obtain a higher concentration of DNA.

• Miniprep were prepared of 2 B. subtilis samples, pUB110 and 18O. They were stored at -20ºC.
• A transformation of 18E will need to be redone; incorrect selection marker was used.

• Ethan is testing the competency of the TOP10 cells he made.

Wednesday, July 31, 2013

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gBlock Fragment Gibson Assembly calculations:

	total fmol (resuspended in 20 ul H2O)	final concentration
gene block 1	648 fmol	.032 pmol/ul
gene block 2	455.4 fmol	.023 pmol/ul

Gibson assembly mix (Trial 1):

• 4 ul gblock 1
• 6 ul gblock 2
• 10 ul master mix

We suspended pub110 insert oligonucleotides in H2O (to 100 uM) and annealed them at 94°C to make duplex dna

Next steps:

• Check the concentration on the oligos against the duplex DNA on the nanodrop to confirm annealing.

August

Week 1

Thursday, August 1, 2013

• Nora’s running PCR for Pveg, 18O, kerA assembly in Kron Lab

Concentration results from nanodrop:

• Forward pUB110 ssDNA: 1596.12 ng/µL
• Reverse pUB110 ssDNA: 1913.96 ng/µL
• Annealed pUB110 dsDNA: 2114.50 ng/µL*

sample	ng/ul	A260	A280	A230	260/230	extinction coefficient
oligo-f	1596	48.4	25.6	22.3	2.17	33
oligo-r	1914	58.0	30.0	26.4	2.2	33
annealed	2159	42.3	22.3	13.8	2.25	50

A = E * b * c

C_annealed = [A / (50 * b)] 2159*(50/33) = 3271

Mass concentration of forward and reverse oligo mixture (ng/ul) = (1596 + 1914)/2 = 1755 ng/ul

• If none of the DNA anneals, the extinction coefficient is 33 and the nanodrop should read 1755*(50/33) = 2659 ng/ul
• If DNA was completely annealed, the extinction coefficient is 50 and the reading on the nanodrop should be 1755 ng/ul

• Therefore, the percentage of annealing is (2659 - 2159)/(2659 - 1755) = 500/905 = 55 %

The assembled DNA was then digested in the following conditions:

• 0.5 ul pUB110 insert
• 0.5 ul NdeI
• 0.5 ul AflII (N)
• 1.5 ul 2.1 buffer
• 12 ul H2O
• 5 ul pUB110 miniprep #1
• 0.5 ul NdeI
• 0.5 ul AflII (N)
• 1.5 ul 2.1 buffer
• 7.5 ul H2O

Friday, August 2, 2013

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• Today we set up digests of pUB110 and 18O.

• Conditions:

• 18O promoter:

• 5 ul 18O miniprep insert
• 0.5 ul EcoRI
• 0.5 ul PstI
• 1.5 ul 2.1 buffer
• 7.5 ul H2O
• Incubate at 37°C beginning at 3 pm

• pUB110 promoter:

• 5 ul pUB110 miniprep insert
• 0.5 ul Afl II (blue-labelled)
• 0.5 ul Nde I (Nora’s)
• 1.5 ul 2.1 buffer
• 7.5 ul H2O
• Incubate at 37°C for 2 hrs

• Yields of pVeg fragment DNA have been low; we will start over by picking more pVeg transformant colonies and cloning more DNA.

Saturday, August 3, 2013

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Sunday, August 4, 2013

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Week 2

Week 3

Tuesday, August 20, 2013

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kerA-pSB1A3 ligation and RFP digestion results

• Measured the concentration of the RFP digested yesterday

• [8/19 RFP Digest] = 74.7 ug/ul

• Ran a 2% gel of the RFP digest

• Lane 2: Ladder (8 ul)
• Lane 4: 8/19 RFP mp#2 Digest (2 ul) + loading dye (2 ul)
• Added 5/10 ul EtBr to the buffer

• There is only 1 band in lane 4.

• Email iGEM to request new Pveg+RBS construct purchased

• Ligated kerA and pSB1A3 with control experiments (Ivan)
• The ligations were set up as follows:

	uncut w/o ligase Control	cut w/ ligase Control	3:1	5:1	10:1
Vol. Insert kerA digest 8/19 (uL)	N/A	N/A	1	1.5	3
Vol. Vector pSB1A3 digest (uL)	(18E)** 1	1	1	1	1
Vol. Buffer (uL)	1	1	1	1	1
Vol. Ligase (uL)	0.5	0.5	0.5	0.5	0.5
Vol. H2O (uL)	8.5	7.5	6.5	6	5.5

used 18E (e. coli promoter from plate 5) instead of pSB1A3, has plasmid [http://parts.igem.org/File:PBca1020-r0040.jpg BBa_J61002] w/ AmpR

• The concentrations of the components of the ligations were as follows:

• kerA 8/19 digest: 90ng/ul
• pSB1A3 digest: 24ng/ul
• 18E miniprep: 963ng/ul

• The ligations incubated at RT overnight
• Another kerA-pSB1A3 ligation is being run in parallel in case the first does not work (Ethan):

• Incubated overnight at 4°C
• ligation reactions:
• ker A gel extraction (8/16) = 115 ng/ul (~1 kb)
• ker A gel extract (8/13) = 75 ng/ul
• pSB1A3 digest = 25 ng/ul (~ 2 kb)

reaction	1	2
kerA	1 ul kerA (8/16) = 115 ng	2 ul kerA (8/13) = 150 ng
pSB1A3	1 ul pSB1A3 = 25 ng	0.5 ul pSB1A3 = 12.5 ng
molar ratio (insert/vector)	2.3	6
	1 ul 10X ligation buffer	1 ul 10X ligation buffer
	0.5 ul ligase	0.5 ul ligase
	6.5 ul water	6 ul water

• Digested (Enzymatic Digestion Protocol)kerA overnight (Ethan)

• 10 ul kerA
• 2 ul digest buffer
• 0.5 ul EcoRI (F.D.)

0.5 ul SpeI (F.D.) 7 ul H2O

• Digested pSB1A3 overnight (Ethan)

• 5 ul pSB1A3 (linear)
• 1 ul digest buffer
• 0.5 ul EcoRI (F.D.)
• 0.5 ul SpeI (F.D.)
• 3 ul H2O

Wednesday, August 21, 2013

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Transformation with kerA-pSB1A3 with control experiments and Transformation of B. subtilis WB700 with pUB110

• Transformed E. coli ([Transformation protocol]) with yesterday’s kerA-pSB1A3 ligation

• used 8/19 kerA digest (90ng/ul), the only pSB1A3 digest (24ng/ul), and 18E miniprep (963ng/ul)
• incubated overnight at 37C starting 2:30pm
• the following experiments, including controls, were performed

plate/tube #	1	2	3	4 (has small crack)	5
tube	3:1	5:1	10:1	cut vector w/ ligase - control	uncut vector w/o ligase - control
Insert (kerA digest 8/19)	1ul	1.5ul	3ul	N/A	N/A
Vector (pSB1A3 digest)	1ul	1ul	1ul	1ul	(18E)** 1ul
Buffer	1ul	1ul	1ul	1ul	1ul
Ligase	0.5ul	0.5ul	0.5ul	0.5ul	0.5ul
H2O	6.5ul	6ul	5.5ul	7.5ul	8.5ul

used 18E (e. coli promoter from plate 5) instead of pSB1A3, has plasmid [http://parts.igem.org/File:PBca1020-r0040.jpg BBa_J61002] w/ AmpR

• About control experiments:

• Uncut vector (BBa_J61002) withOUT ligase: checks for viability of competent cells and antibiotic resistance of the plasmid, as well as transformation procedure itself

• expect colonies
• since we are using a linearized plasmid, is there another uncut vector we could use that has the same antibiotic resistance so we can check the competency of our cells?

• cut vector WITH ligase: shows background due to vector recircularization (and uncut vector)

• expect few colonies (few more than without ligase), since only the plasmids which did not get cut during the digest or the plasmids which did get cut but are able to recircularize with the ligase will be transformed
• this is the most important control. theoretically, there should be no colonies, but there is always some background, so as long as we have a lot more colonies on the vector with insert (actual ligation reaction) plate, it's okay.

• The following controls were not carried out but should be and are useful for future experiments:

• cut vector withOUT ligase: shows background due to uncut vector

• expect few colonies, since only the plasmids which did not get cut during the digest will be transformed
• only do this one if you have extra cut vector, if we don't set it up it's not a big deal, since cut vector with ligase will just show the total background due to uncut vector and vector recircularization.

• insert ONLY WITH ligase: checks for contamination of intact plasmid in ligation or transformation reagents

• expect no colonies, since if there are colonies it indicates contamination of intact plasmid carrying antibiotic resistance in reagents since only the insert was used which should not be able to recircularize into a construct which can be transformed
• this control is optional, but if we are concerned about contamination of reagents with plasmids this could help, so I think we should do it if there is extra insert

• More info on control ligation experiments can be found [http://www.addgene.org/plasmid_protocols/DNA_ligation/ here].

• Analysis of yesterday’s Agarose gel of digested RFP in lane 4:

• the fragment is about 2 kb long
• The BBa_E1010 insert (RFP) is 706bp long, while the vector is 2070 bp long.
• It seems like we have our vector present (the approximately 2kb band in lane 4) but not our insert.

• To check if the BioBrick actually has RFP, plate E. coli with the plasmid with IPTG.
• After checking the kit again, we realized that the BioBrick we isolated from plate 3 well 6G is BBa_R0080, not BBa_E1010.

• The insert in BBa_R0080 is 149 bp, which explains why we couldn’t see it on the gel because it was run off.

• Transformed B. subtilis (Transformation Protocol) with pUB110
  • make O.N. cultures tomorrow in LB Medium

Thursday, August 22, 2013

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Results of kerA-pSB1A3 transformation and control experiments and Transformation of E. coli with RFP

• Transformation Plate Results of kerA-pSB1A3

plate/tube #	1	2	3	4 (has small crack)	5
tube	3:1	5:1	10:1	cut vector w/ ligase - control	uncut vector w/o ligase - control
Insert (kerA digest 8/19)	1ul	1.5ul	3ul	N/A	N/A
Vector (pSB1A3 digest)	1ul	1ul	1ul	1ul	(18E)** 1ul
Buffer	1ul	1ul	1ul	1ul	1ul
Ligase	0.5ul	0.5ul	0.5ul	0.5ul	0.5ul
H2O	6.5ul	6ul	5.5ul	7.5ul	8.5ul
Colonies	>10	>10	>10	0	0*

used 18E (e. coli promoter from plate 5) instead of pSB1A3, has plasmid [http://parts.igem.org/File:PBca1020-r0040.jpg BBa_J61002] w/ AmpR

• The positive control (Tube #5) did not work, even though there should have been colonies. Perhaps there were no colonies because the 18E tube was not actually being the miniprep. The concentration did seem too high.

• Set up three O.N. cultures of kerA-pSB1A3 E. coli in 3 ml LB with ampicillin

• Transformed E. coli with RFP BBa_E1010, well 12N, plate 3 (12N3)

• Followed PCR tube competent cell protocol
• Spread on plates with chloramphenicol and IPTG

Friday, August 23, 2013

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Sending kerA-pSB1A3 Transformants for Sequencing and overnight cultures of RFP transformants

• Isolated Plasmid DNA by Miniprep of 8/18’s O.N. kerA-pSB1A3 cultures #4 and #6 and measured their concentrations to send for sequencing:

colony	ng/ul
4	150 ng/ul
6	185 ng/ul

• The university's DNA sequencing facility recommends that the concentration must be at least size (kb)/10 = [ug/ul]

• 3.35kb / 10 = 0.335ug/ul → 335ng/ul
• The concentrations of the kerA-pSB1A3 above are too low.

• Instead we sent the 3:1 ligation plate (see 8/22/13) for colony picking and sequencing of 5 samples.
• Primers VF2/VR, which are used for sequencing, need to be at 4uM

dilution: (100uM)(2ul)=(4uM)(50ul) ⇒ 2ul primer in 48ul dH2O

• Results of yesterday’s RFP Transformation Plate:

• The colonies are not red. The transformation does not seem to have worked.
• After 5 hours of additional incubation 11am-4pm (20hrs total), white mold started growing on the plate.
• Plate is not cam resistant?

• Transformed E. coli again onto another cam plate, more recently made (agar plates protocol)

• RT SOC incubation 4:50 - 5:50
• Incubation on 37C shaker from 6 pm

• Set up two colonies from yesterday’s RFP transformation plate for overnight culture in 3 mL LB with chloramphenicol to double check if they are cam resistant colonies at 5:30pm.

Week 4

Monday, August 26, 2013

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pUB110 digestions, keratinase assay, and quikchange mutagenesis research

B.subtilis Backbone

• Used FastDigest enzymes and green dye to digest pUB110 for 1 hour
• Ran the pUB110 digest (8/26 pUB110 digest with FastDigest enzymes) on a 1% agarose gel:

• lane 1: ladder (10ul)
• lane 2: fastdigest (20ul)

• The pUB110 digest does not seem to have worked.
• Will try again tomorrow or Wednesday

• Expected sizes:

• uncut pUB110: ~4500 bp
• vector: ~3500 bp
• AflII - NdeI: ~600 bp
• NdeI - NdeI: ~400 bp

B.substilis RFP

• Realized that BBa_E1010 has no promoter so we shouldn’t expect RFP expression anyway.
• Realized that we might not even need the RFP since the instructions for submitting a new biobrick backbone says we only need the prefix/suffix.

Keratinase Assay

• Finished compiling keratin plate/keratinase assay information
• Called Western Live Poultry and Alliance Poultry Farm to ask for chicken feathers. Will be stopping by tomorrow with garbage bags, gloves, and boots to pick up some really dirty feathers that we’ll need to wash before use.

Quikchange Mutagenesis

• For the optimization of keratinase activity
• Quikchange mutagenesis protocols to look at:

1. http://openwetware.org/wiki/Site-directed_mutagenesis
2. http://www.stanford.edu/~loening/protocols/Site_Directed_Mutagenesis.pdf (w/ Pfu)

1. https://www.neb.com/products/e0554-q5-site-directed-mutagenesis-kit
2. https://www.neb.com/protocols/2012/08/29/protocol-for-q5-high-fidelity-2x-master-mix-m0492

• Compiled a protocol for re-using cuvettes.

Tuesday, August 27, 2013

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Troubleshooting pUB110 Digest

• FastDigest enzyme digest may not have worked because we used 1.5/2.1 buffer instead of FastDigest buffer
• We also used too much of the miniprep (5 ul instead of 1.5 ul)when we should have used less.

Wednesday, August 28, 2013

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pUB110 digestion and B. subtilis electroporation

• Bacillus subtilis electroporation: following protocols stated below

• Grew wb700 overnight to OD600 = 4.1 and 7.4
• Started two 10 ml cultures (@ 12 pm) with

• OD600 = 4.1 (500 ul) and
• OD600= 1.3 (750 ul)
• x to OD600 = 0.2 (OD after dilution)

• 4.1 * (0.05) = .205 OD
• 1.3 * (0.075) = 0.0975 OD ???? too low
• subtilis doubling time = 25-40 minutes

• 1:30→ 0.4 abs
• 2:30→ 0.85 abs and 1.0 abs

• Electroporated at 5:30pm, following Eppendorf and its original “High osmolarity improves the electro-transformation efficiency of the gram-positive bacteria Bacillus subtilis and Bacillus licheniformis” (Xue et al. 1999)

• Measured concentration of pBU110 miniprep: [pUB110 miniprep 1] = 367ng/ul sample
• Digested pUB110 overnight with FastDigest enzymes: incubated 6pm

• 1.5 ul pUB110
• 1.5 ul FastDigest buffer
• 0.5 ul AflII
• 0.5 ul NdeI
• 11 ul nuclease-free H2O

• Next Steps:

• Run 1% agarose gel of 8/28 pUB110 FastDigest and 1 ul of undigested miniprep

Thursday, August 29, 2013

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pUB110 digest and Results of B. subtilis electroporation

• 8/28 pUB110 O.N. digest with FastDigest enzymes: incubated 6pm-11:30am

• 1.5ul (miniprep 1: 367ng/ul sample) pUB110
• 1.5ul FastDigest buffer
• 0.5ul AflII
• 0.5ul NdeI
• 11ul nuclease-free H2O

• Ran a 1% agarose gel of 8/28 pUB110 digest (11:45 am - at 120 V)

• lane 1: previously failed digest from 8/27 (5ul dye + 15ul sample)
• lane2: 2 log ladder (10ul)
• lane3: 8/28 pUB110 digest (15ul)
• lane4: miniprep pUB110 (1ul dye + 1ul sample)

• Gel purified (Qiagen gel extraction protocol) the lower two bands: grouped them together into tubes called Upper Band (UB) and Lower Band pUB110 (LB) digest

• Lane 1: Ladder
• Lane 2: 8/28 pUB110 digest

• Measured the concentrations with nanodrop:

• LB: 3.2ng/ul
• UB: 5.2ng/ul

• pUB110 re-digest (with FastDigest conditions, same as 8/26/13)

• used 2ul pUB110 sample with fastdigest conditions, incubated 1hr at 37C (total volume 20ul)
• measured concentrations of digest: 8.1 ng/ul

E.coli kerA Plasmid

• Troubleshooting kerA-pSB1A3 ligations and transformations:

• try using phosphatase to minimize self-ligations
• next digest: scale up 30ul volume, making sure to use water bath in heat block, do it with controls as well (single enzyme and etc)

Friday, August 30, 2013

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kerA-pSB1A3 Sequencing Results

E.colikerAPlasmid

• We recieved our sequencing results of our kerA-pSB1A3 colonies
• When we analyzed the sequences for alignment with our kerA DNA sequence, it was apparent that our kerA biobrick was not ligated into the plasmid.

September

Week 1

Tuesday, September 3, 2013

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Promoter+RFP transformation, kerA-pSB1A3 ligation and digestion, kerA digestion, pSB1A3 digestion

B.substilisRFP

• Transformed E. coli with BBa_J23100 (constitutive promoter family + RFP) for pSB1A3 backbone

• Followed transformation protocol for E. coli PCR tubes
• For making stock pSB1A3 backbone and for trying ligation without gel extraction
• Plates dried in the incubator, but [http://www.protocol-online.org/biology-forums-2/posts/6843.html bubbles at bottom] appeared, which could imply contamination
• Incubation at 37C overnight
• The following morning there were no colonies.
• Must try transformation again except with control experiments.

E.coli kerA Plasmid

• Perhaps we should treat the plasmid with antarctic phosphatase to prevent self-ligation
• Digested kerA and pSB1A3 (Ezymatic Digestion Protocol) overnight at 37°C

Reaction	1	2
	6 ul linearized pSB1A3 backbone = 150 ng	2.5 ul kerA (408 ng/ul) = 800 ng DNA
	1.5 ul buffer	1.5 ul buffer
	0.5 ul EcoRI	0.5 ul EcoRI
	0.5 SpeI	0.5 SpeI
	7.5 ul water	7.5 ul water

• Next Steps:

• kerA-pSB1A3 ligations and transformation of E. coli
• Start O.N. cultures of RFP transformations and cut amp concentration to 50 ug/ul

Wednesday, September 4, 2013

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Results of RFP Transformation, kerA-pSB1A3 ligations and transformation of E. coli, and testing transformation efficiency of E. coli cells

E.coli kerA Plasmid

• Ligated yesterday’s digested kerA and pSB1A3 overnight at 4°C (Ligation Protocol)

• 9 ul kerA digest (40 ng/ul)
• 1 ul pSB1A3 digest (12.5 ng/ul)
• 1.5 ul buffer
• 0.5 ul ligase
• 3 ul water

B.substilis RFP

• BBa_J23100 RFP transformation resulted in no colonies. Must try again.
• Other biobricks with AmpR plasmids we can try transformations with:

• J23119 18A, plate 5 (in pSB1A3)
• J23100 18C, plate 5 re-transform (in BBa_J61002)
• [http://parts.igem.org/Part:BBa_J23101 J23101] 18E, plate 5 (in BBa_J61002)

kerA BioBrick Troubleshooting

• Perhaps, there is something wrong with our kerA?
• Redid kerA biobrick PCR (gibson assembly with geneblocks)

• Measured concentrations of kerA biobrick PCR with nanodrop:

• kerA biobrick PCR #1: 475ng/ul
• kerA biobrick PCR #2: 414ng/ul
• kerA digest 8/21: 50ng/ul

• Tested transformation efficiency of our competent cells with iGEM’s [http://parts.igem.org/Help:Transformation_Efficiency_Kit Transformation Efficiency Kit]: using 5pg/ul, 10pg/ul, 20pg/ul

• Dephosphorylated restriction digests using antarctic phosphorylase:

• 15 ul kerA/pSB1A3 digest (9/3)
• 1.7 ul buffer
• 0.5 ul phosphorylase
• Incubated at 37°C (3:30-4:00 pm)
• Heat inactivated enzyme at 70°C (5 minutes)

• Ligated dephosphorylated pSB1A3 and kerA digests and incubated overnight at 4C

1	2	3
2 ul pSB1A3 Digest (18 ng)	2 ul pSB1A3 Digest (18 ng)	2 ul pSB1A3 Digest (18 ng)
4 ul kerA (188 ng)	8 ul kerA	1 ul buffer
1 ul buffer	1.05 ul buffer	0.5 ul digest
0.5 ul ligase	0.5 ul ligase	6.5 ul H2O
2.5 ul H2O

• Set overnight culture (protocol) of B. subtilis for electroporation tomorrow
• Autoclaved materials and tips and Made more chloramphenicol plates (protocol)

• Transformed kerA-pSB1A3 ligation into DH5Alpha E. coli

• Following PCR tube transformation protocol

1. 1ul DNA into thawed competent cells on ice per tube (3 sets of 3 tubes)
2. Incubate set #1, 2 on ice for 30min, set #3 on ice for 10min
3. Heat shock in PCR machine 45s at 42C
4. Recover on ice for 5min
5. Add to 1mL LB and incubate on shaker for 37C for 1hr
6. Spin down at 8rpm for 30s and plate on Cam plates

Thursday, September 5, 2013

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Results of Transformation efficiency, B. subtilis electroporation (2nd Trial), and RFP transformation

• Results of RFP Control with Transformation Efficiency Kit testing PCR competent cells

• Negative Control Plage

• There weren't any colonies on the negative plate, and we got colonies on the positive plates.
• We can conclude that the competent cells worked.
• However, the number of colonies from plate to plate for the same concentrations weren't consistent. That may be due to inconsistent volume of competent cells in the PCR tubes.

• Electroporation of B. subtilis WB700 (2nd Trial): following "High osmolarity improves the electro-transformation efficiency of the gram-positive bacteria Bacillus subtilis and Bacillus licheniformis" (Xue et al. 1999)

• Calculating Doubling Time Constant

• Average doubling time = 1.156
• ~69min doubling time from 8/28 OD measurements, using the 4.1OD → 0.205OD → 0.85OD or 1OD, taking ave

• Doubling time (formula from wikipedia)

N(t) = 0.85abs C = 0.133abs (see below) d = 1.156 t = 1.156 * [log(.85/.133)/log2] = 3.09 hrs

rough estimate of doubling time to double check above figure:

need for 0.9nm OD

3.46nm/26ml

starting OD calculated at 0.133 (w/o dilution)

x2 = 0.266

x2 = 0.532

x2 = 1.064 ~roughly

O/N culture OD (dilution): 0.346nm (1:10 dilution in water, 100ul in 900ul water) original OD in 15ml tube is 3.46nm

Diluted Initial OD (w/o dilution): 0.133nm (1ml each in 25ml sorbitol and 25ml LB)

Desired OD (w/o dilution): 0.85nm

	0.5M Sorbitol in LB, 25 mL	LB, 25 mL
10am-11:30am	0.133 nm	0.133 nm
11:30 am	0.031*10 = 0.31 nm	0.064*10 = 0.64 nm
12-12:20 pm	N/A	0.104nm*10 = 1.04 nm (used Ethan's spec)
12-1:35pm	0.111nm*10 = 1.11 nm (used Nora's spec)	N/A

LB only

N(t) = 0.85abs

C = 0.64abs (see below)

d = 0.662

t = 0.662 * [log(.85/.64)/log2] = 0.27 *60min = 16min → 20min

[copy, paste into google search] log(.85/.64)/log(2) *0.662

LB 0.5M sorbitol

N(t) = 0.85abs

C = 0.31abs (see below)

d = 1.229

t = 1.229 * [log(.85/.31)/log2] = 1.79 * 60min = 1hr 47min → 1hr 50min

[copy, paste into google search] log(.85/.31)/log(2) *1.229

B. subtilis Electroporation Protocol from Xue et al. 1999:

1. Cool cells in 50ml Falcon tubes in ice-water, 10min
2. Spin down 3,000rpm for 5min (Nora’s lab)
3. Resuspend in 650ul electroporation medium and transfer to 1.5ml tubes
4. Spin down 5,000xg for 5min (BSLC lab)
5. Wash 3x with electroporation medium (500ul)
6. Resuspend in 1/40 of culture volume of electroporation medium

• resuspend in 26/40 = 650ul

• dilution pUB110 #1 244ng/ul: (244ng/ul)(x ul) = (10ng/ul)(200ul) → x = 8.2ul

in 0.1cm gap

Re-using Cuvettes: (protocol) 500ul H2O mix up and down 3x, EtOH mix up and down with 10ul pipette, make sure you see flakes of dead cells on the first mixing, 2x, pour EtOH out as much as possible and air dry for 10min, wipe cap with EtOH too.

number	contains	kV	time constant	notes
1	LBS + DNA	2.0	4.08	no arc
2	LB + DNA	2.0	N/A	didn't wash after initial use, arc'd, immediately filled with EtOH and set aside
3	LBS + DNA	2.1	3.96	new cuvette
4	LBS + DNA	2.0	1.70	incubated sample to see what hapens; washed with 1mL H2O 3x, lots of EtOH 2x, dried out carefully, on plain plate
5	LBS + DNA	2.0	4.16	no arc
6	LB + DNA	2.1	3.06	washed 2x H2O instead of 3x, wasn't as careful, arc'd, incubated
7	LB + DNA	2.1	4.2	washed carefully, no arc
8	LB (N)	2.1	4.2	washed carefully, no arc

Incubated tubes at 6:40pm-9:40pm. Incubated plates at 10pm.

• Plated on 50ug/ml Kan plates → may be issue since protocol recommends 20ug/ml Kan plates?

Friday, September 6, 2013

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Results of B. subtilis electroporation and kerA-pSB1A3 ligations (all trials)

E.coli kerA Plasmid

• Ran a 1% agarose gel of kerA ligations to see if the ligations actually worked

• lane1: ladder (10ul)
• lane2: kerA pcr (0.5ul of 457ng/ul in 5ul)
• lane3: kerA-pSB1A3 ligation 8/15(5ul) [ looks like there some ligation product, but transformation didn’t work ]
• lane4: cut vector w/ ligase 8/20 (8ul)
• lane5: 3:1 ligation 8/20 (8ul)
• lane6: 5:1 ligation 8/20 (8ul)
• lane7: 10:1 ligation 8/20 (8ul)

kerA-pSB1A3 ligation digest 8/20 (3:1 ratio tube)

kerA size: ~1199 bp

pSB1A3 size: ~2155 bp

expected size: = ~3354 bp

B.subtilis Backbone

• Made overnight cultures of B. subtilis BD 366 in 3 mL LB with kanamycin to isolate pUB110 (from 7pm - 1pm)

• B. subtilis Electroporation plates:

• There's only one colony on the plates. It was incubated for longer, and restreaked the following day.

• Transformed (protocol) E. coli with (three) kerA-pSB1A3 ligations and (two) RFP biobrick

• incubated 30 min on ice
• plated on Amp plates

Week 2

Monday, September 9, 2013

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O.N. cultures of past RFP biobricks, kerA-pSB1A3 ligations, and electroporations; Prolonged incubation of electroporations; and Re-streaking RFP biobricks, electroporations, and most recent kerA-pSB1A3 ligation

• At 5pm set up O.N. cultures of:

overnight cultures 9/09

cam: RFP 8/22	overnight culture of a RFP biobrick in pSB1C3 backbone
amp: RFP1 9/6	overnight culture of a RFP biobrick in non-pSB1A3 backbone
amp: RFP2 9/6	overnight culture of a RFP biobrick in pSB1A3 backbone
amp: 10:1 ligation #1	old ligation plates, from 8/20, colony 1
amp: 10:1 ligation #2	old ligation plates, from 8/20, colony 2
amp: 5:1 ligation #1	old ligation plates, from 8/20
amp: 5:1 ligation #2	old ligation plates, from 8/20
amp: 9/6 L2	Ethan’s most recent ligation pSB1A3-kerA, only plate with colonies
kan: AY #3 EP #1	one of my electroporation plates, colony 1
kan: AY #3 EP #2	one of Alice's electroporation plates, colony 2
kan: EB N EP	Ethan’s electroporation plates, negative control
kan: EB 10ng EP #1	Ethan’s electroporation plates, negative control, used 10ng of plasmid DNA, colony 1
kan: EB 10ng EP #2	Ethan’s electroporation plates, negative control, used 10ng of plasmid DNA, colony 2
kan: EB 10ng EP #3	Ethan’s electroporation plates, negative control, used 10ng of plasmid DNA, colony 3
kan: EB 50ng EP #1	Ethan’s electroporation plates, negative control, used 50ng of plasmid DNA, colony 1
kan: EB 50ng EP #2	Ethan’s electroporation plates, negative control, used 50ng of plasmid DNA, colony 2
kan: EB 50ng EP #3	Ethan’s electroporation plates, negative control, used 50ng of plasmid DNA, colony 3
kan: EB 4x EP	Ethan’s electroporation plates, most recent plate that had colonies - did four overnight cultures

• Incubated the following plates longer from 10:30 am and checked again at 4 pm:

plate incubated longer

kan: AY #1 EP	one of Alice’s electroporation plates, wanted to see if more colonies would grow
kan: AY #3 EP	one of Alice’s electroporation plates, wanted to see if more colonies would grow
kan: EB 10ng EP	Ethan’s electroporation plates, wanted to see if more colonies would grow
kan: EB 50ng EP	Ethan’s electroporation plates, wanted to see if more colonies would grow
kan: EB N EP	Ethan’s electroporation plates, wanted to see if more colonies would grow

• Restreaked and incubated until 9pm:

re-streaked

amp: RFP1 9/6	From BBa_J23100, non-pSB1A3 backbone, from Chloe’s RIBS class’ BioBrick used, original plate was messy plating
amp: RFP2 9/6	From BBa_J23119, pSB1A3 backbone, from BioBrick kit, original plate was messy plating
kan: AY #3 EP	plate number 3, my electroporation plate with colonies; replated to try to get more colonies
amp: 9/6 L2	Ethan’s most recent ligation pSB1A3-kerA, only plate with colonies

• The feathers we ordered are here! Must pick them up tonight and we can make our feather keratinase assay.

• Made chloramphenicol plates 250 mL in total (protocol)

• Ligated kerA and antarctic phosphatase treated pSB1A3 and incubated overnight at 4°C (protocol):

1	2	3	4	5
1 ul pSB1A3 Digest (9 ng)	1 ul pSB1A3 digest (9 ng)	1 ul pSB1A3 digest (9 ng)	1 ul pSB1A3 digest (9 ng)	1 ul pSB1A3 digest (9 ng)
0.5 ul ul KerA (220 ng)	8 ul KerA (188 ng)	1.05 ul buffer	1 ul buffer	1 ul buffer
1 ul buffer	1 ul buffer	0.5 ul digest	0.5 ul digest	0.5 ul digest
0.5 ul ligase	0.5 ul ligase	6.5 ul h2o	6.5 ul h2o	6.5 ul h2o
2.5 ul h2o total DNA = 22 ng/ul

Tuesday, September 10, 2013

Wednesday, September 11, 2013

Thursday, September 12, 2013

Friday, September 13, 2013

Saturday, September 14, 2013

Week 3

Monday, September 16, 2013