Notebook

5/30/2022

Reporter Options:

• beta-galactose (colorimetric)
• luciferase (light)
• RFP (fluorescence)
• pH reporter assay
• ELISA (colorimetric)

If not our microplate reader, how will we detect color change?

Magnetic bacteria: We need to domesticate more microbes

Homework:

• Casey to look through 3 previous iGEM projects
• Everyone to search/be ready to describe 2-3 options for reporter assays
• Beth to ask Chaz to look into Arduino/Raspberry Pi sensor options

Notes on labwork:

Goal: get a feel for 3A cloning/assembly

• *biobricks (see 3A Assembly Protocol)
• Goal: get a feel for 3A cloning/assembly
• Standard set of restriction enzymes --> swap out parts like legos
• Used Pst I and EcoRI
• Anything under an iGEM project should be 3A assembly compatible
• Everything has the same "standard connection" from an engineering point of view
• You can stitch things together at different restriction sites

In our plasmid we have PstI and EcoRI on either side of the insert (RFP or GFP) --> flanked by the restriction enzymes

Depending on which 3A backbone plasmid (different antibiotic resistance) --> have to be outside of the section GFP/RFP flanked by the Pst1 and EcoR1:

• Amp
• Kan
• Chlor

Cut & digest with PstI and EcoRI:

GFP/RFP (writing for GFP but same idea)

Physically cutting out the GFP but the pSB1A3 (AmpR) backbone is the same in both GFP and RFP --> we ligate the opposite GFP and RFP into the opposite backbones --> it works because the EcoR1 and Pst1 will still attach at the same sites --> all will grow on Amp plates

Ligation means to "glue back together"

Last step: transformation --> transfer the new plasmids into E. coli onto Amp plates

Those that have succesfuly been transferred into E. coli survive / the E. coli without the plasmids transferred will die because they lack Amp resistance

7/1/2022

The start of getting the actual sequences for this paper: 10.1074/jbc.272.52.32824

Essentially this was an attempt to start rebuilding some of the coding regions that were talked about in that paper, even though I'm not sure I know what I'm doing The first thing I did was find the FASTAs for human AhR and ARNT, as well as a two way promoter that promotes in both directions when exposed to galactose and is repressed by glucose. The paper then has this image:

And so took the reverse compliment of the ARNT dna and stuck it all together end to end, which is the ARNT rev comp> GAL 1,10 2way promoter > AhR sequence. Unless there's something I'm missing, in theory this base sequence should directly code for the genes in the diagram.

This is all I had time for at this current time due to my program ending and a flurry of final project, but now that I will be going back to New York soon I will be dedicating a lot more time to this project and should have draft plasmids ready to go fairly quickly.

8/13/2022

Notes from BT re: Gibson Assembly

Blog Post - takeaway messages:

• Recommends 15-40bp overlaps with melting temp > 48C
• Look out for secondary structure in the overhang sequence
• Works best with fragments > 200bp (exonuclease may digest fragment before it anneals/polymerizes)

8/30/2022

• Measured and mixed 8g LB agar and 200 mL and autoclaved at 121 C for 20 min.
• After autoclaving, added 200 uL of 50 mg/mL Ampicillin Stock(1000x)
• Poured 15-20 mL per plate
• Dried the plates for 30-35 minutes
• Streaked tonight

Yeast Plasmids Parts

• AHR / ARNT sensor (13kb) with URA3 selection marker
• CYPC1A / Luciferase reporter (11kb) with LEU2 selection marker (CYPC1A is eukaroyote promoter)
• Backbone BF303B
• Gene inserts Twist (6 pieces, each piece 1800 bp) for 5.8kb

Use Gibson assembly to stitch backbone and Twist inserts to make PLK001

Notes

• ori for bacteria and 2ori for yeast
• Grow plasmids in bacteria to make more plasmids

Things to Validate/Test

• Response of CYPC1A to AHR / ARNT
• Quantity of luciferase to PCB

8/31/2022

• Transferred 20mL of LB Broth to a 50 mL conical tube in the sterile hood
• Added 20 uL of 50 mg/mL Ampicillin Stock (1000x) to the conical tube in the sterile hood
• Transferred 3 mL each of LB + Amp into six 15 mL conical tubes
• Transferred single colonies from plates grown overnight to tubes
• Three tubes were inoculated with a single colony each of pBF3038
• Three tubes were incolulated with a single colony each of pIGAhRC
• Rotated at 37 C overnight

9/1/2022

MiniPrep

• Followed MiniPrep according to Genspace Protocol(https://benchling.com/s/prt-jvYx7x8SrS9lZZC9TITZ?m=slm-LFfwpQR2WMXVxY4j3DaJ), started with 1.5 mL of culture.
• On the last step, eluted with 30 uL Elute Buffer, centrifuged 1 min., picked up flow through, reapplied flow through the column, and centrifuged for 1 min.

QuBit

• Followed Genspace Protocol
• Used 2 uL of plasmid DNA Sample and 198 uL Mastermix

Results:

	Luke	Naomi	Maya
pBF3038	157 ug/mL	192 ug/mL	184 ug/mL
pIGAhRC	58.7 ug/mL	68.9 ug/mL	64.5 ug/mL

Glycerol Stock:

• Made a 30% Glycerol Stock
• Used 500 mL of 60% Autoclaved Glycerol and 500 mL of Culture
• Stored -80C Freezer

Reconstituting Primers:

• Took brand new primers, labeled them with shorthand names, and diluted them to 100 uM in the original container by adding 10x of their molecular weight in sterile water
• Made a working stock(10 uM) by adding 90 uL of sterile water, 10 uL of diluted sample, vortexed.
• Stored in -20C FreezerTher

9/7/2022

Day 1: Bacterial Transformation

• Take SOC media out of the fridge to warm to Room Temp (use the one that's labeled expired 6/21, not Open Plant's bottle) - take a 650ul aliquot out in the sterile hood
• Take 3 LB+Amp plates out of the fridge to warm to Room Temp
• Thaw 4 aliquots of cells on ice
  • Gibson: Add 5ul of chilled assembly product to competent cells (mix gently by flicking, do not vortex)
  • Positive Control: Add 1ul of the pBF3038 diluted 1:10 in water
  • Positive Non-Diluted Control: Add 1ul of the pBF3038
  • Negative Control: no DNA
• Place the mixture on ice 30 min
• Heat shock @42C for 30 seconds
• Place mixture back on ice for 5 min
• Add 200 ul of SOC to the tube
• Add 250 ul of cells + media to the LB+Amp plates - spread using the glass beads
• Incubate O/N @37C

9/8/2022

Day 2: Check Results + Start O/N cultures for Miniprep

Take plates out of the incubator (AM), store at 4C until ready to use

At ~5-8pm

• Take LB out of the fridge, aliquot 3ml each for 2-6 clones (depends on how efficient the transformation is) in 15ml Conical tubes + Amp
• so for 2 clones, 6ml LB + 6ul Amp --> mix and split into 2 tubes
• 6 clones = 18ml LB + 18ul Amp --> mix and split into 6 tubes
• Make sure lids are on tight, and place in the end-over-end rotator inside the 37C incubator (turn on the rotator)
• Grow O/N

9/9/2022

Day 3: Miniprep & Qubit

Follow Miniprep Protocol and Qubit as before

• Miniprep Protocol followed exactly (added 30 ul of Elution Buffer)
• 4 minipreps were created with the four colonies that were rotated overnight
• 4 Qubit samples from the 4 minipreps

Used 3 ml of overnight rotations in the miniprep

G1	G2	G3	G4
34.3 ug/ml	16.4 ug/ml	20.8 ug/ml	8.95 ug/ml

Qubit samples were slightly vortexed to ensure accurate results. Qubit was recalibrated

Sample 1	Sample 2	Sample 3	Sample 4
21.9 ug/ml	10.9 ug/ml	26.1 ug/ml	11.0 ug/ml

Day TBD: Sequencing

Genspace Protocol (+ get some assistance from Jehovani or arrange a Zoom call with BT to complete these steps)

Modifications to the standard PCR product ordering:

Sample Prep

• Prep Samples for Sequencing with same labeling: 1 on the lid, AA01 on the side, etc. in strip cap PCR tubes
• • Aim for 10-15ul at 50ng/ul
• Prep Primers in 500uL total volume at 5uM in 1.5ml tubes (25ul of 100uM stocks + 475 fancy sterile nuclease-free water)

Azenta Website (ask Jehovani/BT for help)

• Select Plasmid
• Select Predefined (may use premixed depending on the number of clones you want to sequence)
• Select Standard Priority
• Enter the # of samples
• Order Name: iGEM2022 pLK001 Clones
• DNA Name: pLK001_# of the clone
• Length: <500bp
• Concentration: Include from Qubit (and/or standardize concentrations to 50ng/ul) and
• Primer Name: Include all primers: 1F;1R;2F;2R;3F;3R;4F;4R;5F;5R;6F;6R;7F;7R
• Submit

Shipping/FedEx - follow the protocol (ask Jehovani/BT for help)

Day TBD: Yeast Transformation

Making Media: Yeast SDO Plates -Ura AND -Leu

Genspace Media Protocol, adapted recipe:

• 20g/L Bacteriological Agar
• 6.7g/L Yeast Nitrogen Base
• 1.62g/L Yeast Synthetic Drop-out Medium -His, -Leu, -Trp, -Ura
• 0.4g/L L-Tryptophan
• 0.2g/L L-Histidine

+ make Dextrose/glucose spearately - recombine after autoclaving, before pouring plates (as with other YPD Agar)

*Also make a batch with no agar for liquid cultures

Yeast Transformation

*Grow fresh cells from BY4741 stocks

Select transformants to work with - make frozen glycerol stocks

Assay Function

Restriction digest of pBF3038 to remove the 1500bp sequence between the f1 ori and CYC1 Terminator (make the plasmid compatible with the Gibson Assembly Twist fragments)

• XbaI (CYC1)
• SpeI-HF (f1 ori)

NobCloner

Reagent	25uL Rxn
500 ng DNA	2.5
10x CutSmart	2.5
XbaI	0.5
SpeI-HF	0.5
Nuclease-free H2O	19
	25

• Digest at 37C for 15 min
• Run 0.7% agaroe gel, 1x TAE, 2.5ul EtBR
• Run gel for ~30 min @120V

5500 bp fragment

Gel Extraction Protocol

• 336mg gel + 4x vol (1344ul) melt @ 52C for ~10 min
• Elute in 7ul nuclease-free water

Qubit Protocol

• Digested, purified pBF3038 - 8.8ng/ul
• Proceed to Gibson Assembly

Gibson Assembly

• Digested pBF3038 to 1500bp remove fragment and make space for our Gibson construct
• Conducted gel electrophoriesis to seperate plasmid fragments and purify product
• Qubit(https://benchling.com/s/prt-krVZlD6784pKdJ29HRo6?m=slm-VPeDJdGsdan633YoKcuU) to measure DNA Concentration (8.8ng/ul)

	Weight (ng)	Vol to 10ng/ul	Length (bp)	pmol	Volume (ul)	total pmol for Gibson
CYP1A1_1	765	76.5	1149	0.0133895695	1.3	0.0174064404
CYP1A1_2	588	58.8	1142	0.0134716422	1.3	0.0175131349
CYP1A1_3.1	757	75.7	871	0.0176631635	1.1	0.0194294798
CYP1A1_3.2	828	82.8	599	0.025683832	1	0.025683832
CYC1	675	67.5	377	0.0408079984	0.7	0.0285655989
Akaluc CDS	847	84.7	1705	0.0090232348	1.9	0.0171441462
pBF3038		5	5500	0.0024615385	5	0.0123076923
TOTAL					12.3	0.1380503244

Gibson Assembly Protocol

Component	7 Fragment Assembly Vol (ul)
CYP1A1_1	1.3
CYP1A1_2	1.3
CYP1A1_3.1	1.1
CYP1A1_3.2	1
CYC1	0.7
Akaluc CDS	1.9
pBF3038	5
Gibson 2x MM	25
Nuclease-free H2O	12.7
Total Volume	50

Notes:

• Due to low concentration of digested plasmid, we opted to go for ~equimolar ratios vs. total pmol
• Due to >6 fragments assembled, we opted for 25ul instead of the recommended 10ul 2x Gibson Master Mix
• We also let the reaction go for 90 min (instead of 60)
• BT went ahead and did a transformation on 9/9 just to test... (may need to order backup supplies and that will take some time to ship)
• Rest of the Gibson Assembly product stored at -20C until the team can continue with the transformation

9/16/2022

BOOMS

• 1X TAE running low -> made from 20mL of 50X TAE + 980mL dH2O
• Spe-HF running low -> spun it down in centrifuge
• Split 1 digest sample into 2 gel wells -> extraction results in low quatity (2ng/ul)
• Qubit mixed up BR and HS standards -> redid standards while plasmids waited for a bit -> inaccurate results (measured the second time after some time and concentration was drastically different, e.g. Luke's sample was originally 2.08ng/ul and became >20ng/uL to next few times measured after some time elapsed)
• Autoclave LB agar used parafilm (melting) + Amp resistance left out a bit longer before adding to LB agar
• Negative control plates used dirty beads -> redid only the steps for heat shock

1. Make Gel

   Use 6-well comb, make 0.7% agarose gel (50uL TAE + 0.35g agarose), 1x TAE, 2.5ul EtBR

2. Disgest Plasmid pBF3038

   Add the following reagent into a PCR tube:

   	Luke	Naomi	Maya
   pBF3038	157 ug/mL	192 ug/mL	184 ug/mL
   pIGAhRC	58.7 ug/mL	68.9 ug/mL	64.5 ug/mL

   1000ng = 1ug, 184ug per 1000uL = 184ng/uL

   Reagent	50uL Rxn	50uL Rxn
   1000 ng DNA	5.4347826087	5.4
   10x CutSmart	5	5
   XbaI	1	1
   SpeI-HF	1	1
   Nuclease-free H2O	37.5652173913	37.6
   Total Volume	50	50

   Digest at 37C for 15 min

3. Make Media for Plating

   Measured and mixed 8g LB agar and 200 mL of distilled water and autoclaved at 121 C for 20 min (3rd conical flask settings). After autoclaving, added 200 uL of 50 mg/mL Ampicillin Stock (1000x). Poured 15-20 mL per plate. Dried the plates for 30-35 minutes.

4. Run Gel

   • 15uL of 100kb extended DNA ladder
   • Add 10uL of dye into each 50uL sample (1:6X loading dye)
   • Load 50uL into 2 wells each (should give 4 times as much)
   • Load very slowly, will spill if too fast
   • Run gel for ~30 min @120V
   • Loaded into 3 wells

5. Gel Extraction

   Follow protocol from Monarch DNA Gel Extraction Kit

   • Tube 1: 0.19g ; Tube 2: 0.39g (split tube into 2 smaller ones)
   • Tube 1: 190mg (760ul)
   • Tube 2A: 0.182g (728ul)
   • Tube 2B: 0.211g (844ul)
   • use 2 spin columns
   • 8uL of elution buffer
   • combine the 2 spin column volumes (2uL for Qubit) => 12uL
   • reload elution buffer for a second spin

6. Qubit

   Aim for concentration of digested, purified pBF3038 > 20ng/ul (5uL = 100ng)

   • Tube 1: 20.1 ug/ml
   • Tube 2: 2.08ug/ml

7. Gibson Assembly

   Protocol reference from NEB

   • 50–100 ng of vector with 2-3 fold molar excess of each insert
   • Can aim for 200ng of vector
   • pmol = (concentration*1000)/(D8*650)
   • Available stock: 200uL 2X Gibson MM
   • Do 2-3 Gibson reactions to test:
   • Tweak pmols of inserts
   • Tweak more Gibson MM or total volume to make sure insert fragments less than 20% (~40uL)
   • Incubate samples in a thermocycler at 50°C for 90 minutes
   • Store samples on ice or at –20°C for subsequent transformation.

   	Weight (ng)	Vol to 10ng/ul	Length (bp)	pmol	Volume (ul)	total pmol for Gibson
   CYP1A1_1	765	76.5	1149	0.0133895695	2.1	0.028118096
   CYP1A1_2	588	58.8	1142	0.0134716422	2.1	0.0282904486
   CYP1A1_3.1	757	75.7	871	0.0176631635	1.7	0.0300273779
   CYP1A1_3.2	828	82.8	599	0.025683832	1.2	0.0308205984
   CYC1	675	67.5	377	0.0408079984	0.8	0.0326463987
   Akaluc CDS	847	84.7	1705	0.0090232348	3.2	0.0288743515
   pBF3038		6	5500	0.0056223776	4.9	0.0275496503
   TOTAL					16	0.2063269214

   Component	7 Fragment Assembly Vol (ul)
   CYP1A1_1	2.1
   CYP1A1_2	2.1
   CYP1A1_3.1	1.7
   CYP1A1_3.2	1.2
   CYC1	0.8
   Akaluc CDS	3.2
   pBF3038	4.9
   Gibson 2x MM	17.5	half of total volume
   Nuclease-free H2O	1.5
   	35

8. Plating

   • Take SOC media out of the fridge to warm to Room Temp (use the one that's labeled expired 6/21, not Open Plant's bottle) - take a 650ul aliquot out in the sterile hood
   • Take 3 LB+Amp plates out of the fridge to warm to Room Temp
   • Thaw 3 aliquots of cells on ice
   • Gibson: Add 5ul of chilled assembly product to competent cells (mix gently by flicking, do not vortex)
   • Positive Control: Add 1ul of the pBF3038 diluted 1:10 in water
   • Negative Control: no DNA
   • Place the mixture on ice 30 min
   • Heat shock @42C for 30 seconds
   • Place mixture back on ice for 5 min
   • Add 200 ul of SOC to the tube
   • Add 250 ul of cells + media to the LB+Amp plates - spread using the glass beads
   • Negative: Just competent cells
   • Positive: pBF dilution
   • Incubate O/N @37C

10/1/2022

Materials:

• 4 Erlenmeyer flasks
• Weigh boats
• G and ml scale

Yeast Inoculation:

• Poured 11.5 mL of YPD Broth into a conical tube with approximatgely 50 uL of stock S. cerevisciae
• Incubated in rotator at 30C for 5 hours --> BY 4741
• Custom Media for selecting for our plasmids:
• Values in table obtained from Sigma Aldrich
• Started with:
• SDO --> syntetic droupout media (SDO-4)
• his
• trp
• leu
• ura
• Histodine and Tryptophan get added back to end up the desired missing components
• his
• trp
• leu
• ura

We need 10 plates with 20 mL per plate so 20 mL per plate x 10 plates = 200 mL (for Agar)

Sigma Aldrich's concentrations are in mg/L or g/L, so we took 1/5 of all of those values to find out how much we needed, displayed in the table below.

Flask 1	Concentration (mg/L)		Flask 2	Concentration (mg/L)		Flask 3	Concentration (g/L)		Flask 4	Concentration (g/L)
Trp	76	15.2	Trp	76	15.2	Dextrose	20	4	Dextrose	20	4
His	76	15.2	His	76	15.2	Bact Agar	20	4	YNB	6.7	1.34
SDO-4	1390	278	SDO-4	1390	278

Flasks were autoclaved at 121 degrees C / 26 minutes after each component was added to the flasks, listed above

Continuously quantified the Optical Density of the Yeast inoculation as it was in a rotator

• Added 1 mL of growing yeast culture and empty media to cuvettes and compared their optical densities
• 0.02 around 11:00
• 0.1 around 2:30
• 0.18 around 4:30

Poured plates, only added dextrose to plates since the liquid media should be kept seperate of galactose (in agarose) or glucose so as not to disrupt the GAL1,10 promoter. Plates will have both galactose and glucose, so they should cancel each other out (hopefully).

Flask 5:

• Created because dual expression vector does not express if there is glucose but expresses if there is galactose --> this flask contains a concentration of galactose ONLY --> for the controls
• 0.016% D-Galactose g/100 ml --> 0.032 g/200 ml (making 200 ml total in flask)
• still adding 32 mg to 100 ml water because we will add flask 5 into flask 2 --> need ratios to be the same (ratios of things we put in them to be the same) --> might not add flasks in entireties
• Autoclaved flask 5 for 26 minutes at 121 degrees C

5/21/2022

Notes For The Bacterial Degredation of Polychlorinated Benyphyls

• This procedure is taking genes from prokaryotic bacteria to create eukaryotic yeast
• The goal is to combine a total of 9 genes and insert them into 5 plasmids from 4 different bacteria into yeast
• The previous team was only able to do 1 or 2 genes successfully
• BphAI+2 was one of the genes that successfully assembled as DNA
• Only the BPHA1 protein was able to be verified (opperating correctly and forming mRNA). It was verified with Western blot using Flag-tag.
• The PCBA5 enzyme was successfully applied to transfer the PCBs from highly chlorinated to less chlorinated
• Five enzymes are needed to be applied to make the PCBs n-chlorobenzoate with 2-hydroxy pentanoate
• Three enzymes are then needed to be applied to make the benzoate pyruvate to acetyl coa

5/28/2022

• Measured concentration by Nanodrop
• Calculate volume of plasmid and amount of water needed to get 25 uL of each template

Template DNA	Concentration (ng/uL)	A260/280	Volume of Plasmid (uL)	Amount of Water (uL)
RFP	11.1	1.83	2.3	22.7
Joy	103.4	2.07	0.24	24.76
AFP Plasmid	16.7	1.85	1.5	23.5
11.2 Plasmid	17.1	1.8	1.46	23.54

7/30/2022

Performed ligation and transformation of original PCBA5 gene. No successful bacteria colonies.

8/6/2022

Repeated transformation with 3 samples:

• Positive control (RFP; J04450 plasmid)
• PCBA5 gene
• Negative control (water)

Plated on LB chlor plates and incubated at 37 C

Transformation:

• Thaw 3 tubes 25uL competent cells on ice.

*Dry off the lid and label them Lig (1 for each student), RFP, GFP, and Ctrl (for Negative Controls)

• Add 1uL of ligated DNA to Lig tube and mix gently by flicking the tube 4–5 times. Do not vortex. Add 1uL of 5 ng/ul diluted plasmid plasmid to the corresponding positive control tubes. No DNA into Ctrl tube.
• Place the mixtures on ice for 15 minutes. Do not mix.
• Heat shock at 42°C for 30 seconds. Do not mix.
• Place on ice for 5 minutes. Do not mix.
• Add 200 µl of room temperature LB media to the tube.
• Label the bottom of your LB Agar plates with the Sample ID, Your Initials, and Date
• Spread 150 µl of the cells and plasmid mixture onto the appropriate plates using the glass beads.

*Dispose of used beads in the dirty bead container (do not throw away)

• Incubate overnight at 37°C.
• Analyze results during the next session.

8/20/2022

Resuspending, digesting, and ligating PCBA4, PCBA1, and PCBA5 with sticky-end revised to the vector PSB1c3.

PCBA1	24.4 ng/uL	0.88
PCBA4	29.2 ng/uL	0.80
PCBA5	25.0 ng/uL	0.96
PSB1c3	21.4 ng/uL	0.94

8/27/2022

• Transformed bacteria following protocol
• Plated bacteria

9/10/2022

• Attempt to culture bacteria from 8/27 (ie. with sticky-end linear vector) yeilded no successful colonies
• Set up following 10 uL ligation reaction with PCR products (PCBA1, PCBA4, PCBA5) in separate tubes:

Reagent	Volume	Final Concentration
5x Ligase Buffer	2 uL	1x
Blunt-end linear vector	1 uL	20 ng/uL
PCR product	1 uL	30 ng/uL
Molecular Biology-Grade Water	5 uL
T4 DNA Ligase	1 uL	200 U/uL

• Incubated ligation reaction at 25 C

9/17/2022

• Rhodocuccus medium synthesis:
• Stock salt solucion: Missing MnSO4, left out HCl from solution
• Concentrated Goodies requires Stock Salf, which is incomplete as of 9/17
• Mg/Ca?B1 goodies mix: everything except Concentrated Goodies mixed together and stored at 4 C
• Basal Salt solution: complete
• Blunt-end ligation in E. coli
• Transform bacteria following protocol (30 min before heat shock, 30 sec heat shock, hour incubation after SOC addition)
• Plate bacteria in LB Growth plates
• Check on growth next Saturday

9/24/2022

Screening E. coli colonies

9/17 bacteria (SmartJoin plasmid)

• Ligation negative control = 0 colonies
• Negative coltrol = 0 colonies
• Ligation positive control = 30 colonies
• PCBA1 = 1 colony
• PCBA4 = 1 colony
• PCBA5 = 12 colonies

8/27 bacteria (PSB1c3 plasmid)

• Negative = 0 colonies
• Positive = 3 pink, 6 white
• PCBA1 = 2 pink, TMTC white
• PCBA4 = 1 pink, TMTC white
• PCBA5 = 3 pink, 1 white
• PCBA1 Lig = 10 pink, 7 white
• PCBA4 Lig = 30 pink, 22 white
• PCBA5 Lig = 38 pink, 25 white

Performed colony PCR on the plated colonies

Performed gel electrophoresis on the amplified DNA for screening of PCBA1, PCBA4, and PCBA5

Completed synthesis of Rhodocuccus growth media, 100 mL total

10/1/2022

Rhodococcus Growth Media Procedure

• Create the stock salt solution by combining 22.94 g/L MgCl2-6H2O, 2.0 g/L CaCO3, 4.5 g/L FeSO4-7H2O, 1.44 g/L ZnSO4-7H2O, 0.85 g/L MnSO4-H2O, 0.25 g/L CuSO4 - 5H2O, 0.24 g/L CoCl2-6H2O, 0.06 g/L H3BO3, 51.3 mL HCl
• Create the concentrated goodies by combining 50 mL stock salt solution, 3.009 grams of MgSO4, 25 mL 1% FeSO4
• Bring the volume of the concetrated goodies up to 100 mL
• Sterilize by filtration and store in fridge
• Create 100X Mg/Ca/B1/Goodies mix by combining 60 mL 1M MgSO4, 3 mL 1M CaCl2, 3 mL 10 mM thiamine, and 75 mL concentrated goodies
• Adjust volume to 300 mL, sterilize by filtration and store at 4 degrees
• Create basal salts by combining 30 gram KH2PO4, 60 grams Na2HPO4, 5 grams NaCl, 10 grams NH4Cl
• Autoclave to serilize
• Create final growth media by combining 100 mL 20% dextrose, 10 mL Mg/Ca/Bl/Goodies, 200 mL basal salts, and 690 mL water

Rhodococcus Optical Density Procedure

• Measure optical density of starting cell cultures (Rhodococcus, E. coli)
• Calculate the total volume of the cell cultures c1 v1 = c2 V2
• Create Rhodococcus growth media by mixing 100ml 20% dextrose, 10ml Mg/Ca/Bl/Goodies, 200Ml basal salts, and 690 ml water
• Combine Rhodococcus and Rhodococcus growth media in a flask
• Combine E. coli and Rhodococcus growth media in a flask
• Combine E. coli and LB growth media in a flask
• Combine Rhodococcus and LB growth media in a flask
• Place all flasks into the incubator shaker and check the concentration of cells every half hour for 5 hours

Time	Rhodocuccus in Rhodocuccus media	Rhodococcus in LB media	E. coli in Rhodococcus media	E. coli in LB media
Initial	0.306	0.306	0.630	0.630
30 mins	0.064	0.099	0.138	0.172
60 mins	0.063	0.087	0.150	0.214
90 mins	0.065	0.080	0.166	0.276
120 mins	0.064	0.065	0.190	0.394
150 mins	0.064	0.210	0.067	0.631
180 mins	0.120	0.103	0.238	0.891
210 mins	0.085	0.109	0.344	1.149
240 mins	0.081	0.098	0.481	1.500
270 mins	0.102	0.113	0.634	1.731
300 mins	0.106	0.094	0.776	1.856

10/8/2022

Repeating last week's experiment at 30 C instead of 37 C

Time	Rhodocuccus in Rhodocuccus media	Rhodococcus in LB media	E. coli in Rhodococcus media	E. coli in LB media
Initial	0.191	0.191	2.850	2.850
30 mins	0.054	0.086	0.093	0.117
60 mins	0.039	0.063	0.048	0.096
90 mins	0.067	0.066	0.131	0.104
120 mins	0.055	0.073	0.051	0.138
150 mins	0.060	0.090	0.058	0.220
180 mins	0.077	0.081	0.072	0.267
210 mins	0.074	0.135	0.088	0.330
240 mins	0.068	0.072	0.115	0.425
270 mins	0.104	0.101	0.123	0.484
300 mins	0.185	0.126	0.109	0.570
350 mins	0.171	0.180	0.140	0.721
380 mins	0.170	0.200	0.123	0.763
410 mins	0.187	0.217	0.128	0.851
470 mins	0.256	0.327	0.137	1.127
520 mins	0.364	0.432	0.156	1.303