iGEM 2019 — Team Sydney_Australia

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# Team:Sydney Australia/Notebook

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* Team

  [The Team](https://2019.igem.org/Team%3ASydney_Australia/Team)
   [Collaboration](https://2019.igem.org/Team%3ASydney_Australia/Collaborations)
* Project

  [Description](https://2019.igem.org/Team%3ASydney_Australia/Description)
   [Design](https://2019.igem.org/Team%3ASydney_Australia/Design)
   [Experiments](https://2019.igem.org/Team%3ASydney_Australia/Experiments)
   [Notebook](https://2019.igem.org/Team%3ASydney_Australia/Notebook)
   [Characterisation](https://2019.igem.org/Team%3ASydney_Australia/Contribution)
   [Results](https://2019.igem.org/Team%3ASydney_Australia/Results)
   [Demonstrate](https://2019.igem.org/Team%3ASydney_Australia/Demonstrate)
   [Improve](https://2019.igem.org/Team%3ASydney_Australia/Improve)
   [Attributions](https://2019.igem.org/Team%3ASydney_Australia/Attributions)
* Parts

  [Parts Overview](https://2019.igem.org/Team%3ASydney_Australia/Parts)
   [Basic Parts](https://2019.igem.org/Team%3ASydney_Australia/Basic_Part)
   [Part Collection](https://2019.igem.org/Team%3ASydney_Australia/Part_Collection)
* [Safety](https://2019.igem.org/Team%3ASydney_Australia/Safety)
* Human Practices

  [Human Practices](https://2019.igem.org/Team%3ASydney_Australia/Human_Practices)
   [Outreach & Engagement](https://2019.igem.org/Team%3ASydney_Australia/Public_Engagement)
* Achievements

  [Medals](https://2019.igem.org/Team%3ASydney_Australia/Medals)
  [Modelling](https://2019.igem.org/Team%3ASydney_Australia/Model)
  [Software](https://2019.igem.org/Team%3ASydney_Australia/Software)

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Notebook

We've been working hard inside and outside of the lab for almost five months now, and whilst it has been
challenging we've had a lot of fun! Read more about it below.

Week 1: 20/5/19 - 26/5/19

* 23/5/19 - Team meeting

First ever team meeting - the project begins! So exciting!

Week 2: 27/5/19 - 2/6/19

* 31/5/19 - Team meeting

Team roles decided.

Week 3: 3/6/19 - 9/6/19

* 7/6/19 - Wet lab

Safety induction for the team - learning how to be safe in the lab

* 7/6/19 - Team meeting

Planning wet lab work, outreach measures, and human practices initiatives

Week 4: 10/6/19 - 16/6/19

* 14/6/19 - Team meeting

Discussed current lab work, further planning on outreach, discussed the need for Human Ethics approval for us to conduct the desired interviews for HP, discussed logo.

Week 5: 17/6/19 - 23/6/19

* 19/6/19 - Wet lab

Fahad prepared an overnight culture of E. coli containing pk18, pET28 and PUS250 plasmids.

Week 6: 24/6/19 - 30/6/19

* 24/6/19 - Wet lab

Fahad transformated pET28 and pk18 into TOP10 E. coli* 25/06/19 - Wet lab

Fahad and Benj made a culture of pET28, pk18 and PUS 250. pET28 and pk18 were pelleted and stored in the -20C freezer. PUS250 had not reached a high enough concentration so was left to incubate overnight.

Merrie and Isobel made an overnight culture of Top10 E. coli on LB agar in incubator at 37 degrees overnight for preparation of competent cells.

* 26/6/19- Wet lab

Merrie made up a 10mM Tris PH8 from 1M stock solution and sent for autoclaving. Merrie also pelleted the overnight culture of PUS250. The 1:10 dilution had OD600 of 0.502.

Merrie completed plasmid DNA extraction of pET28, pk18 and PUS250 plasmids according to the Coleman lab protocol. The only variation to the protocol was that the pellets were resuspended in 1XTE pH8 instead of 10mM Tris as there was no sterile Tris. All plasmids were stored in the -20 freezer.

Isobel and Fahad performed a restriction digest of Pet28 PUS250 & PK18 with HindIII and EcoRI. The restriction digest was conducted according to manufacturer's protocol for ThermoScientific FastDigest restriction enzymes, and products were run on a 1% agarose gel in 1xTAE for 30 minutes at 200 V. The gel was run to quantitate and identify the prepared plasmids.

Isobel and Fahad performed a restriction digest of Pet28 PUS250 & PK18 with HindIII and EcoRI. The restriction digest was conducted according to manufacturer's protocol for ThermoScientific FastDigest restriction enzymes, and products were run on a 1% agarose gel in 1xTAE for 30 minutes at 200 V. The gel was run to quantitate and identify the prepared plasmids.

pK18 and pET28c(+) were grown in a 50 mL overnight culture of LB, at 37°C with shaking at 200 rpm. 5 x 10 mL of each culture was decanted into separate falcon tubes, then spun at 2500 rcf for 15 minutes. Supernatant was discarded and 500 uL of P1 resuspension buffer was added to each tube. After resuspending the pellet by vortex, 2 x 250 uL of each tube was added to separate microcentrifuge tubes, and miniprep proceeded according to manufacturer's protocol (ISOLATE II Plasmid Mini Kit, Bioline).

10 x 50 uL of purified plasmid was obtained for pK18 and pET28c(+), and samples for respective plasmids were combined into single tubes. Plasmid concentration was determined by spectrophotometer (ThermoFisher NanoDrop 2000c), and were found to be 281.6 ng/μL for pK18 and 38.1 ng/μL for pET28c(+).

To confirm, a restriction digest was conducted according to manufacturer's protocol for ThermoScientific FastDigest restriction enzyme, and products were run on a 1% agarose gel in 1xTAE for 30 minutes at 200 V. The results were consistent with the nanodrop.

* 27/6/19 - Wet lab

Merrie performed a digest of the Vivid G blocks. Each reaction (WT, CH1, CH2, CH3, CH4) was incubated for 15min at 37ºC then run through spin column to remove enzymes and eluted in EB buffer (15μL).

Merrie performed a digest of the PET28 plasmid. The reaction was incubated for 15min at 37ºC then run through spin column to remove enzymes and eluted in EB buffer (45μL).

Merrie performed a ligation reaction of the PK18 with each of the Vivid GBlocks (WT, CH1, CH2, CH3, CH4). The reactions were incubated at room temperature for 15 minutes and stored in -30 freezer overnight.

Isobel continued with process of making TOP10 competent cells by taking 2x 3mL cells from overnight incubation into and putting them into 100mL LB broth.

Initial OD600 ~0.1. Incubated in 37ºC shaking 200rpm until OD600 ~0.5 reached. The cells were pelleted at OD600 ~0.45 as per protocol - the pellet was hard to resuspend. Cells were divided into 70 220μL aliquots stored in -80ºC freezer.
28/06/19-Wet lab-Isobel, Benj, and Merrie performed a heat shock transformation of TOP10 Ecoli with pk18 Vivid ligation. Positive and negative controls were generated. Transformed cells were spread-plated onto LB-Km-Xgal plates and incubated at 37C overnight.

Fahad performed a PET28 purification using spin column using Coleman lab protocol and eluted in 20μL EB buffer. Final concentration was 38.1ng/μL.

Fahad also made up a stock of LB-Xgal-Km plates.

* 29/6/19 - Wet lab

Fahad performed a plate observation for the transformed Vivid cells. No successful transformations were observed.

Week 7: 1/7/19 - 7/7/19

* 2/7/19 - Wet lab

Isobel and Benj performed a Vivid gene digestion and ligation into PK18. The plasmids and G-blocks were digested using EcoRI and HindIII for one hour at 37C. The enzymes were then heat killed at 80C for 20mins. 10uL plasmid digest and 10uL Vivid digest were combined with 10uL digest buffer and 1uL T4 ligase and incubated for one hour at room temp.

Isobel and Benj then performed a heat shock transformation of ligated PK18 and Vivid G-blocks. Positive, negative, ligation, and digestion controls were generated. 100uL of the transformed cells were spread-plated onto LB-Km-Xgal plates. The plates were incubated at 37C overnight.

* 3/7/19 - Wet lab

Checked plates. Very few colonies observed, almost all were blue. White colonies did not fluoresce under UV light so presumed false positives. Decided to test a new ligation protocol the next day.
4/7/19-Wet lab/outreach prep-Isobel conducted a plasmid preparation for the WMBB activity. Plasmids B29, PUS116 & PK18 to be used for fruit genetics experiment. Each plasmid was grown inside TOP10 E. coli with the appropriate antibiotic in 1mL LB until it reached OD600 1.5. Each culture was then transferred to a 50mL LB broth to grow at 37C until it reached OD600 1.0. Each culture was then mini-prepped according to Coleman lab protocol.

Isobel also prepared part of the fruit/plasmid mixture for the WMBB activity. For further details please see our [WMBB write up.](https://2019.igem.org/Team%3ASydney_Australia/Public_Engagement)* 4/7/19 - Wet lab

Merrie and Benj performed digest and ligation testing of PK18 to troubleshoot the Vivid transformations. Learned to decrease plasmid concentrations for future cloning. Confirmed that ligation was working as intended.

* 5/7/19 - Wet lab

Benj performed another digestion and ligation control test. Plasmid digested with various enzymes was run on a gel to ensure that the plasmid was being cut as expected. The band sizes and numbers were consistent with the digestions working correctly. Plates were prepared with cells transformed with either digested or digested+ligated plasmids. Significant differences were seen in between the colony counts of the digested and ligated plates, indicating a significant improvement in digestion when the concentration of plasmid was reduced for digestion.

* 5/7/19 - Wet lab/outreach prep

Nathan continued preparing for the WMBB outreach activity by mixing the watermelon supernatant with the plasmid mixtures. For further details please see our [WMBB write up.](https://2019.igem.org/Team%3ASydney_Australia/Public_Engagement)

* 5/7/19 - Team meeting

Discussed current Vivid progress, finalised logo, ironed out details for WMBB outreach event, discussed contacting Usona - major psilocybin producer for clinical trials.

* 6/7/19 - Wet lab/Outreach prep

Isobel and Nathan continued to prepare for the WMBB outreach event by performing a restriction digest on B29, PK18, and PUS116 and running a gel with the Kiwi DNA and the plasmids. For further details please see our [WMBB write up.](https://2019.igem.org/Team%3ASydney_Australia/Public_Engagement)

Week 8: 8/7/19 - 14/7/19

* 8/7/19 - Wet lab/Outreach prep

Isobel ran a gel with the three different fruit DNAs and the three different plasmids. She found that as the strawberry + pk18 produced the clearest band pattern that would be the designated ‘sick person’ sample for the activity.

She also decided that for the students all their reagents would be replaced with water and then the gel replaced so the students would experience the experimental procedure without the risk of not getting easily interpretable results. See our [WMBB write up.](https://2019.igem.org/Team%3ASydney_Australia/Public_Engagement) for more details

* 8/7/19 - Wet lab

Benj formulated a revised cloning protocol for his vivid genes. This created Vivid Cloning Protocol Version 1. See our [protocols section](https://2019.igem.org/Team%3ASydney_Australia/Experiments) for the full protocol. He performed this new protocol on his Vivid genes.

* 9/7/19 - WMBB outreach event

Nathan and Isobel ran a short molecular biology class with Indigenous youth called ‘Fruit Forensics’ where students had to use molecular biology to find out which piece of fruit had given their patient food poisoning. See our full [WMBB write up.](https://2019.igem.org/Team%3ASydney_Australia/Public_Engagement) for more details!

* 9/7/19 - Wet lab

Benj had great success today! Finally saw some positive colonies on his CH2 and CH3 plates. However, unfortunately there were still no positive colonies for WT, CH1, and CH4. These vivid gene blocks needed to be re-done using Vivid Protocol Version 2.

* 10/7/19 - Wet lab

Benj performed Vivid Protocol Version 2 (see our protocol section for full protocol - link) on the WT, CH1, and CH4 gene blocks. Due to a plate mix-up the newly transformed cells were not plated onto differential media so there was some delay in seeing results.

* 11/7/19-Wet lab

Benj had even more success! He got some positive patches on his plate. There was some contaminating blue colonies among the white patches so these were further purified by streak plating.

* 12/7/19 - Wet lab

Nathan performed some Psi gene cloning of PsiM/D/K into PET28. He got positive colonies for all three genes.

Week 9: 15/7/19 - 21/7/19

* 15/7/19 - Wet lab

Benj and Isobel worked on purifying the Vivid colonies. Unfortunately several blue colonies remained on the plates. Benj consulted Nick (supervisor) who suggested that the constitutive expression of PK18 may be stressing the cells. New patch plates were incubated at 30C to reduce the stress on the cells.

Isobel was able to streak out 2x CH3 white colonies from CH3-1 and CH3-2 plates from 11/7. She then streaked CH2 from patch plate square 3 from 12/7. Lastly she made a patch plate of CH1 colonies from 10/7 pellet plate with 3x controls and 6x white colonies.

Benj then patch plated WT, CH4 from spread plates made on 10/7. CH2 was patch plated from spread plates from 8/7. CH1 was streak plated from CH1 patch plates from 12/7 using squares 1, 2, 4 and 5. All the plates that were made up were incubated at 30°C to reduce stress on the cells.

Lastly, Benj and Isobel made 19 LB-KM-XGAL plates (400mL LB agar, 500μL Kanamycin, 1mL X-Gal) and 19 (LB-KM plates 400mL LB agar, 500μL Kanamycin).

* 16/7/19 - Wet lab

Benj continued the purification of the Vivid transformations. He made streak plates from the patch plates made on 15/7 of WT and CH4 onto LB-Km-Xgal plates.

* 17/7/19 - Wet lab

Benj found that the plates grown at 30°C showed distinct and isolated white colonies on all Vivid streak plates.

He re-streaked the white colonies onto LB-km-Xgal plates and KM-charcoal plates. The charcoal plates reduce fluorescence of LB and therefore background fluorescence, good for testing the fluorescence of the different harmonisations. Benj also performed repurification on KM-Xgal plates of CH1, CH2, and CH3, then made some more KM-LB-Xgal plates.

Benj and Nathan performed colony PCR on the Psi D, K, and M plates. Four positive colonies were obtained for PsiM, three positive colonies for PsiK, and seven positive colonies were found for PsiD. However, whilst bands were present some of the dye had migrated off the gel resulting in poor visualisation of the ladder. Benj and Nathan decided to run a gel again using three positives for each Psi gene to get better visualisation of the ladder and bands for each gene.
18/7/19-Wet lab-Benj and Nathan ran a gel using three positive colonies per Psi gene from yesterday. This gel confirmed their results. These positive colonies were streak-plated on LB-Km plates and inoculated into LB-Km broths, all cultures were incubated at 36C overnight.

* 19/7/19 - Wet lab

Benj and Nathan removed the Psi gene cultures from the incubator. The plates were stored in the 4C fridge. The cultures were transferred to 5mL falcon tubes and the cells pelleted. Excess supernatant was removed and the tubes were placed in the -30C freezer for 24hours.

Benj found that the CH1 AND CH3 cultures have stable incorporation of the Vivid/pk18 plasmid but CH2 and CH4 and WT appear to be rejecting the plasmid this may be due to the high copy number of the plasmid and strong expression of the VIVID protein. To test this theory he grew new plates of CH1 and CH3 and incubated them at 30C and 37C to test whether the E. coli would reject the plasmid at different temperatures.

* 20/7/19-Wet lab

Benj and Nathan removed the frozen tubes from the freezer and thawed them. Any remaining supernatant was removed with careful pipetting. The cells in each tube were resuspended in 1mL EB buffer before being transferred to individual 1.5mL eppendorf tubes. Plasmid was then extracted from tubes using the 'Plasmid miniprep protocol (5 ml culture) – Spin Column Method' protocol, except the tubes were left to dry at room temperature on a clean Kimwipe for 10mins instead of at 60C to evaporate residual ethanol. All plasmid samples, eluted into 30uL of 10mM Tris-HCl pH8, were then placed in a fridge at 4oC for 48 hours.

Benj purified and moved the CH1 and CH3 plates from 19/7/19 to the cool room.

Week 10: 22/7/19 - 28/7/19

* 22/7/19 - Wet lab

Benj checked in on his Vivid colonies. Found that the CH1 plates were stable at 37C and 30C whilst the CH3 plates were beginning to turn blue at 37C but still stable at 30C.

Both the CH3 30C and 37C plates were moved to 30C to confirm it is the speed of growth that is resulting in the reversion to blue colonies.

WT, CH4, and CH2 were re-streaked on Km-Xgal and grown at 30C.

* 23/7/19 - Wet lab

Nathan analysed the PET28-PsiK/M/D plasmids prepared on 20/7/19 for purity and concentration using a nanodrop spectrophotometer. The least contaminated tube for each Psi gene was selected for restriction digest preparation.

A single restriction digest using EcoR1 and a double restriction digest using EcoR1 and HindIII was then set up for each plasmid sample as well as an empty pET28 vector, all tubes were incubated for one hour at 36C.
15uL of each sample was then mixed with 2uL of 6X purple loading dye before being pipetted into individual wells on an Agarose gel for gel electrophoresis.

Nathan found that the results indicated that the PsiK and PsiM colonies selected were suitable for protein purification study. Unfortunately, PsiD showed double banding - indicating the presence of both an empty pET28 and the PsiD construct within the cells. Using these cells for protein purification study would result in suboptimal expression of the PsiD gene. Nathan decided to test the other tubes using restriction digest before performing protein purification steps for PsiD. New individual 5mL of LB/Kan broths were inoculated with the tested PsiK and PsiM colonies.

Benj cultured Vivid CH1 and CH3 in 5mL Km-LB overnight to prepare for mini-prep. More colonies from the WT-patch plate were streaked out and incubated at 30C. The CH2 and CH4 plates from 22/7 were re-streaked and incubated at 30C.
24/7/19-Wet lab-Benj plasmid purified the CH1 and CH3 cultures according to the Coleman labs nanodrop protocol. CH1 had a concentration of 434.6ng/uL and CH3 had a concentration of 30.7ng/uL.

* 25/7/19 - Wet lab

Nathan performed another restriction digest analysis on the other two tubes of PsiD cells. Results revealed no double banding in the single digest column for tubes 2 and 3, with a distinct size difference when compared to the empty pET28 plasmid. This meant that tubes 2 and 3 were suitable for protein purification work.

Nathan also performed PCR on the PsiH gene block to add BamH1 & Xba1 restriction sites to the construct to make it suitable for insertion into the PCW plasmid.

Benj performed another mini-prep on the CH1 and CH3 cells according to Coleman lab protocols. The concentrations were similar to 24/7: CH1 had a concentration of 431.3 ng/uL, CH3 had a concentration of 28ng/uL. He then performed a restriction digest on both of them and ran similar amounts of plasmid on a gel along with pk18 cut and uncut plasmid. No double banding was observed. It does appear that the CH3 plasmid is significantly lower in concentration than the CH1 plasmid in E. coli Top10 cells, possibly due to the constitutive expression stressing the cells.

Benj decided to try transforming the vivid genes into PUS250 - an inducible plasmid. Benj used the leftover G-blocks for CH2 and CH3. For CH1 and CH3, the positive colonies from prior experiments were mini-prepped and that DNA was digested then used to transform into PUS250.

* 26/7/19 - Wet lab

Benj grew the CH2 AND CH3 spread plates for 3 days at 37C. White colonies were patch plated on KM-Xgal plates and KM-Cumate.

The LB plates were used to determine which colonies were truly fluorescent. Cumate (the inducer) plates should fluoresce strongly if Vivid has been successfully transformed into the pus250 plasmid and into the E. coli
Benj also patch plated white colonies from the digestion and ligation plates to compare expression of CH1 and CH3 to the white colonies.

* 26/7/19 - Team meeting

Discussed current Vivid progress, discussed Psi-gene successes, made a plan for putting PsiH in PCW plasmid. Discussed going to the SBA Conference, discussed Dusseldorf postcard collaboration.

Week 11: 29/7/19 - 4/8/19

* 30/7/19 - Wet lab

Isobel performed a nanodrop on the PCW plasmid stocks in the lab. Found that the stock had a concentration of 825ng/uL. Diluted the stock 1 in 4. Performed two double digests and two single digests using NdeI and SalI. Unfortunately, the gel had no discernable bands - possible due to the plasmid being degraded in the cool room, enzymes being heat killed, or the mung bean endonuclease that was added to blunt the ends not being in the appropriate concentration.

Fahad prepared an overnight culture of the PUS250 plasmid using two colonies from 100uL heat shock transformed plate were isolated and to incubate 2x 5mL vials of LB-Km. They were incubated in the shaking 37C incubator overnight.

* 31/7/19 - Wet lab

Isobel and Merrie transformed some TOP10 E. coli with the remaining PCW (as per Coleman lab protocol) as very little stock remained after the failed digestion. Grew 100uL aliquots of the supernatant and pellet of the transformed cells on Ampicillin plates, plus a TOP10 only negative control.

Merrie used 1 sample of the overnight PUS250 culture for miniprep of plasmid DNA. Miniprep of 5mL sample as per Coleman lab protocol with no changes. Nanodrop showed 18ng/uL DNA.

* 1/8/19 - Wet lab

Isobel and Merrie found that the supernatant and pellet plates from 31/7 had good growth and there was no growth on the negative control plate. A big loop-full of cells taken from pellet plate, vortexed in 1mL broth, then inoculated into 50mL LB with Ampicillin. Grown in 37C shaker for 24hrs.

* 2/8/19 - Wet lab

Merrie miniprepped the PCW cells from 1/8 and tested the concentration using the nanodrop. The concentration was 113ng/uL. A restriction digest was then performed on the PCW plasmid DNA to remove the p450 from the plasmid using Sal1 and Nde1.

The double digest, single digest, and uncut PCW plasmids were run on a gel. The use of LAB buffer in the gel resulted in poor resolution in the ladder making it hard to determine size. However, the double digest had a double band, indicating the removal of p450, which made the cut PCW plasmids suitable for end-blunting and ligation.
The ends were blunted using mung-bean nuclease and the DNA purified using a spin-column. The yield was low as spin columns do not work well on DNA over 5kb. The ends were ligated together using T4 ligase.

Benj found that the transformation of of CH2 and CH3 in PUS250 into TOP10 appeared to have failed. Fluorescence did not vary between transformed and untransformed colonies. The CH2 & CH3 patch plated on LB-Km-Xgal and LB-Km-Cumate had similar fluorescence and growth to digestion and ligation controls. Benj also performed a restriction digest on the various CH1-PK18 cultures from previous weeks.

* 2/8/19-Team meeting

In depth human practices meeting, got updated on the process of our Human Ethics approval - still waiting. Decided our main focuses are ethics, medical, and economics. Also discussed our outreach theme and potential for iGEM to be integrated into the high school curriculum.

Week 12: 5/8/19 - 11/8/19

* 5/8/19 - Wet lab

Isobel tested the concentration of the digested and ligated PCW plasmid using the nanodrop. The concentration in one replicate was 569.9ng/uL, and in the second replicate 712.5ng/uL.
She then ran the plasmids on a gel to check for a band of the correct size. The gel made it apparent how much yield had been lost from the column purification, a phenol chloroform purification would have been better.

* 6/8/19 - Wet lab

Nick performed error prone PCR done using primers NVC15b and 16b to amplify the VVD-36-CH4-C37A gene (CH4 codon variant) out of pK18 plasmid backbone, from a colony of E. coli TOP10 carrying pK18-VVD-CH4 plasmid. PCR as normal using Taq polymerase, except 0.15 mM MnCl2 added to make it error-prone. Band looks good on gel.

* 7/8/19 - Wet lab

Isobel ran PCR on the PCW plasmid to test whether the digested plasmid still contains p450. The primers were designed to span the region where p450 used to be. If the p450 was cut out, end up with 700bp product. If p450 still in, end up with approx 2000bp product.

* 7/8/19 to 8/8/19 - JAMS Science Week Booth Outreach Event

Isobel, Merrie, Nathan, Emma, and Benj volunteered to help JAMS Sydney with their Science Week booth at the Australian Museum. They helped kids with activities such as looking at pond water under a microscope, viewing their own skin or hair using a handheld microscope, and viewing plates containing different microbes. See our full [JAMS write up](https://2019.igem.org/Team%3ASydney_Australia/Public_Engagement) for more details!

* 8/8/19 - Wet lab

Isobel ran a gel of the PCR product from 7/8. Found that the PCR product band was around 700bp, indicating that our PCW plasmid had its p450 successfully removed - ready to be replaced by PsiH!

* 9/8/19 - Wet lab

Nick purified the error-prone PCR product from 6/8 on column, digested with EcoRI+HindIII, the mixture was heat-killed, then ligated to pK18 that had been digested with same enzymes overnight at 4C.

* 9/8/19 - Team meeting and meeting with Alex Kelly

Talked to Alex Kelly, biotechnology entrepreneur and iGEM Sydney alumni, about the considerations we must think of regarding scaling up our project for industry.

* 10/8/19 - Wet lab

Isobel performed a PCR on the PCW plasmid with p450 inside to double check that the PCW plasmid tested on 8/8 had its p450 removed and it wasn’t a PCR error. She ran the whole-PCW PCR product on the same gel as the digested-PCW PCR product and found that the band sizes were as expected.

Nick transformed the error-prone PCR ligation mix into TOP10 cells, which were spread on several LB-Km-Xgal plates at different dilutions, then incubated 30C for 48 h.

* 11/8/19 - Wet lab

Nick examined the error-prone PCR plates under white light and with long-wave UV lamp. Under white light, approx. 50% appear recombinant (white colonies), while rest are blue (vector only). Under UV lamp, a mix of non-fluorescent and weakly fluorescent colonies seen, but also a few colonies showing obviously brighter fluorescence – these were picked and restreaked for later analysis (total 12 colonies).

Week 13: 12/8/19 - 18/8/19

* 12/8/19 - Wet lab

Isobel performed a colony PCR on colonies transformed with the digested-PCW plasmid to ensure that the colonies used for mini-prep and insertion of PsiH were p450-free plasmids.
The PCR-product from the colony PCR was run on a gel. Found that all seven colonies tested had the correct PCW plasmid in them. Isobel selected one colony to grow up for mini-prepping.

* 12/8/19 - Human Practices

Emma finished writing up the literature review on the History of Psilocybin in Society. Read it on our  [Human Practices page!](https://2019.igem.org/Team%3ASydney_Australia/Human_Practices)

* 13/8/19 - Wet lab

Emma, Nathan, and Fahad performed a miniprep of the PsiK and PsiD PET28 plasmids to prepare for sequencing.

The mini-prep was performed according to protocol with minor alterations: all samples were spun down into two columns instead of four, and the samples were eluted with only 12uL EB to increase the concentration of the final yield.
The final concentration of PsiD was 119.3ng/uL. The final concentration of PsiK was 71.7ng/uL.

Merrie performed a PCR to add Bam1 and XBa1 restriction sites to PsiH. However, unfortunately this PCR was unsuccessful.

* 14/8/19 - Wet lab

Isobel performed a miniprep of pCW-no-p450 and then digested with BamH1 HF and Xba1. In the plasmid with the p450 still in, there's two restriction sites and the fragments would be approx 7900 and 800bp. In the plasmid with no p450, there's only 1 restriction site for BamH1 and fragment would be 7000bp.

The restricted pCW-no-p450 on the gel, there was only single bands in the BamH1 digest lane, so could presume that p450 had been removed.

Nathan grew more E. coli TOP10 containing PET28 PsiD/K in 50mL LB/Km, 50ug/mL, 37C overnight.

Fahad made more protein extraction buffer C made according to Coleman lab protocol. pH buffered to 8.5.

Benj prepared a 5ml culture of Vivid wild-type in Km-LB and ch1-5 to miniprep for sequencing.

Merrie re-tried the PCR to add Bam1 and XBa1 restriction sites to PsiH. The PCR product was run on a gel. There were nice bands between 2kB and 3kB as expected. The bands were cut out, weighed, and moved to the cool room for purification later.

* 14/8/19 - Team meeting

Outreach focused meeting: discussed the upcoming innovation games outreach event. Made a shopping list of supplies, plan for activities and posters, and decided who would be coming on the day.

* 15/8/19 - Wet lab

Nathan pelleted and froze the PsiD/K PET28 cells were cells at -30C with supernatant removed.

Benj performed a miniprep on his WT, CH1, and CH3 cells to prepare for sequencing. He tested the concentration of DNA using the Nanodrop. Final concentrations for WT, CH1, and CH3 (tubes 1 and 2) were 21ng/uL, 357ng/uL, 64.2ng/uL and 51.4ng/uL respectively.

Benj also performed some further purifications on Vivid CH1, and streak plated more colonies of CH4.

Merrie performed a gel DNA extraction on the PsiH from 14/8. The final concentration according to the nanodrop was 8.3ng/uL - not ideal!

* 15/8/19 - Human Practices

Our human ethics approval came through!

* 16/8/19 - Wet lab

Nathan performed a PsiD/K-PET28 plasmid purification using 800uL P1 added to defrosted cell pellets. This was split between 2x 4 epindorff tubes (200uL in each) before the 5mL Coleman purification protocol was followed.

Benj performed a 5mL miniprep on two CH4 colonies, CH2, and CH3.

* 16/8/19 - Human Practices

Emma completed her literature review “The What and How of Psilocybin”. Read it on our  [Human Practices page!](https://2019.igem.org/Team%3ASydney_Australia/Human_Practices)

* 18/8/19 - Innovation games outreach event

Went to Sydney Olympic Park for the outreach event. Isobel, Merrie, Nathan, Benj, and Emma were there on the day. Fahad and Nathan made posters. Nathan and Benj ran a fluorescent protein activity where children could look at E. coli art. Merrie ran a strawberry DNA extraction demonstration. Emma ran a DNA modelling activity using paper and lollies. Isobel ran a creative activity where children could either draw a scientist or draw a hybrid animals using randomised ‘genes’. See our full [Innovation Games write up](https://2019.igem.org/Team%3ASydney_Australia/Public_Engagement)for more details!

Week 14: 19/8/19 - 25/8/19

* 20/8/19 - Human Practices

First round of emails to experts inviting them to be interviewed was sent out by Nick Coleman to the experts on Emma’s list.

* 20/8/19 - Wet lab

Merrie and Isobel attempted to perform a gel purification on the PsiH PCR product to prepare for PUS381 ligation. The first attempt had low yield (8.3ng/uL), the second attempt had a much better yield (24ng/uL).

* 21/8/19 - Wet lab

Merrie performed the PsiH/PUS381 ligation. Transformed TOP10 E. coli with the ligation product (as well as a positive control of PUS381 and a negative control of no DNA) using the Heat Shock protocol.

* 21/8/19 - Team Meeting

Discussed current Vivid optimisation work versus error-prone PCR work. Discussed Psi progress. Told Emma to stop being addicted to making her Human Practices lit reviews. Celebrated that we finally have Human Ethics approval and can contact our HP experts and organise interviews! Discussed USYD Open day outreach event, who would be there, and what we will do. Booked our flights to Boston!

* 21/8/19 - Human Practices

First two interview subjects responded positively to being interviewed. Emma sent emails to arrange one phone interview and one email interview.

* 22/8/19 - Wet lab

Merrie did patch plating and colony junction PCR of 24 colonies.
Isobel ran the PCR products on 2x 1% TAE (50mL) with 7.5uL GelGreen. 100V, 40min. Loaded 5uL PCR product and ladder.
Only one positive colony was found. Isobel streaked out a plate from that colony and inoculated a 50mL broth for use in a miniprep. The PsiH+PUS381 plasmid was named PUS382.

* 23/8/19 - Human Practices

Emma completed her Safety of Psilocybin literature review. Read it on our  [Human Practices page!](https://2019.igem.org/Team%3ASydney_Australia/Human_Practices)

* 23/8/19 - Wet lab

Nathan received sequencing results back for PsiD/K/M. Decided PsiD needs resequencing close to the centre of the gene as there were many apparent deletions and low confidence regions.
The PsiM construct was sequenced well with no deletions or substitutions. The PsiM-Pet28 construct can now be renamed PUS383.

PsiK had a few areas of low confidence sequence that need to be resequenced.

* 24/8/19 - Wet lab

Benj prepared patch plates of the different Vivid harmonisations.

Week 15: 26/8/19 - 1/9/19

* 26/8/19 - Wet lab

Nathan attempted to elute protein extracts of PsiD/K/M from TOP10 cells according to the protein extraction protocol. He ran these extracts on a gel and found no protein bands on his SDS-PAGE gel.

Benj streaked out more vivid colonies from patch plates made on 24/8 onto individual streak plates for the different Vivid harmonisations.

Nick amplified the inserts from his more-fluorescent error-prone-PCR colonies with primers NVC15b /16b, the PCR products were then purified with column, and sent for Sanger sequencing with primer 15b.

* 27/8/19 - Wet lab

Merrie performed a restriction digest of PUS381 using HindIII, ScaI, EcoRV, and Xba. Ran restriction products on 0.8% agarose TAE 100V 45 mins with 5uL gel green.
Fahad performed a restriction digest on PUS382 using HindIII, ScaI, and Xba. Ran on a 0.8% 1xTAE agarose gel 50mL at 100V 60 min.

* 28/8/19 - Wet lab

Isobel redid the restriction digests of PUS381 that Merrie performed the day before as the resulting bands were too diffuse. Found that the bands were the correct size.

Nathan redid the PsiD/K/M protein extraction according to the protein purification protocol from the TOP10 cells containing the respective plasmids. Again, he got no product on his gel. He discovered that TOP10 cells were not appropriate for making the His-tag purification proteins. He checked the original workflow and found that this work was meant to be performed in BL21 cells.

Benj restreaked individual colonies from plates prepared on 26/8 to further purify.

Merrie set up an indole assay using PCW as a positive control and PUS382 as a test. Loops full of each plate’s colonies were transferred to 10mL LB-Carb and incubated, shaking, till the OD increased by 0.5. IPTG and Alanine was added to both cultures and left shaking overnight.

* 28/8/19 - Team meeting

Discussed progress of the codon harmonisation program. Benj was directed to prepare for the Vivid fluorescence assays, Nathan discussed his difficulties with protein purification, Merrie discussed her indole assay progress. Proposed that the next meeting needs to be a wiki working bee to get more of our writing on track.

* 29/8/19 - Wet lab

Nathan transformed TOP10 cells with PUS381 using the heat shock protocol. Incubated at 37C overnight on LB-Amp plates with a negative control.

Benj prepared 5mL individual cultures of each of the Vivid harmonisations to prepare for mini-prep. The vivid harmonisations were also plated onto charcoal plates to evaluate fluorescence.

The results of the indole assay from 28/8 were observed. A strong rust/purple colour change was observed in both. Decided to repeat the experiment with negative controls to be sure of results.

* 30/8/19-Wet lab

Isobel inoculated 2x 50mL LB-Carb broths with large loopfuls of PUS381 transformed cells from the supernatant plate Nathan prepared the day before.

Benj mini-prepped the 5mL cultures he made the day before.

Emma prepared some negative controls for the indole assay conducted on 28/8/19. PUS381 (no p450 or PsiH) induced/uninduced and PUS382 uninduced. Loops full of each plate’s colonies were transferred to 10mL LB-Carb and incubated, shaking, till the OD increased by 0.5. IPTG was added to one of the PUS381 cultures, alanine was added to all, then all were left shaking overnight.

* 31/8/19 - Wet lab

Benj noted the results of the negative controls indole assay - rust/purple colour change was seen in all cultures. This was unexpected and it was decided that the experiment needed to be repeated again. P.S. Little did they know, this experiment would continue to trip them up for many weeks to come!

* 1/9/19 - Wet lab

Benj performed a plasmid digest on several of his Vivid constructs. Some of the cells containing the tested constructs were incubated overnight in 5mL cultures to get enough DNA to confirm the band size, while other tested cells were streaked out on solid media for further tests.

Nick analysed the error-prone PCR sequences from 26/8 by aligning them to original VVD-CH4 sequence in Snapgene. 10/12 sequences gave good reads over the whole plasmid insert. many point mutations were seen in these clones, but all 10 mutants that had complete sequences shared a mutation at methionine-130, either M-->I or M-->V or M-->L. A new Gblock of VVD-CH4 was designed and ordered, which contained only the mutation M130I, to test if this was solely responsible for the increased fluorescence. This part would henceforth be referred to as VVD-Met-130, after the position in the DNA sequence that was mutated to generate the extra-fluorescent phenotype.

Week 16: 2/9/19 - 8/9/19

* 2/9/19 - Wet lab

Isobel miniprepped PUS381 then performed restriction digests on PCW, PUS382, and PUS381 to ensure that the plasmids used in the indole assay were correct. All plasmids’ digests were looking distinct from one another.

* 3/9/19 - Human Practices

Received interview response from Dr Francisco Moreno. Read our ‘Psilocybin Talks’ [here!](https://2019.igem.org/Team%3ASydney_Australia/Human_Practices)

* 3/9/19 - Wet lab

Merrie prepared PsiD and K for sequencing. She also prepared the PUS382 plasmid for sequencing.

Nathan performed PCR on the PsiD plasmid to add the BSA1 restriction site for our Golden Gate Cloning procedure. When the products were ran on gel there was some background banding, so gel purification was required.

Nathan also prepared overnight cultures of PsiD/K/M in both non-inducing and auto-inducing media. 50mL media in sterile 250mL bottles with two loopfuls of transformed BL21 cells.

Merrie and Emma transformed TOP10 cells with PCW, PUS382, and PUS381. The freshly transformed cells were transferred straight into the 10mL LB-Carb cultures. Cultures failed to grow.

* 4/9/19 - Wet lab

Benj set up a PCR of his tested Vivid recombinant constructs.

Nathan began protein extraction on the cells that were grown overnight. Extraction was performed using the Protein Extraction protocol. Protein concentration was estimated using the Nanodrop spectrophotometer. ~40ug each was run on an SDS page gel. Finished gel was stored in the cool room.

Nathan got sequencing results back for PsiD which revealed three deletions that would render the protein non-functional. He went into the cool room and found older plates of PsiD cultures from 17/7 and sent the other PsiD+ colonies off for sequencing after performing a junction PCR to amplify the region.

Merrie and Emma re-attempted the direct transformation to indole assay. Once again, the cultures failed to reach an appropriate OD. This was due to insufficient concentrations of cells - turns out there’s a reason why you plate your transformed cells first!

* 4/9/19 - Team meeting

Wiki working bee! Discussed the wiki progress at length and assigned wiki writing tasks. Had a lot of fun!

* 5/9/19-Wet lab

Isobel performed spanning and junction PCRs of PCW, PUS382, and PUS381 to confirm the results of her restriction digest from 2/9.

Benj performed more PCR on his Vivid constructs to amplify the region for sequencing.

Merrie transformed more TOP10 cells with PCW, PUS381, and PUS382. Once complete, they were plated on LB-Amp plates and incubated overnight.

* 8/9/19 - Wet lab

Isobel freshly transformed some TOP10 cells with PCW, PUS382, and PUS381. Benj streaked out some PUS381 to prepare for a colony PCR. Spanning PCR gave normal results, PsiH specific PCR gave weird results - possible contamination of our PsiH primer.

Week 17: 9/9/19 - 15/9/19

* 9/9/19 - Wet lab

Benj performed a colony PCR on the pure Vivid isolates using 2x Mango mix. Used three cultures per harmonisation type, plus one PK18 control and one no template control. He also restreaked several of the Vivid isolates onto Km only plates as the Xgal appeared to be interfering with the fluorescence of colonies. He also prepared some 5mL cultures of several Vivid isolates to incubate overnight.

Nathan performed a colony PCR on the PsiD colonies picked from the 17/7 plate.

Isobel performed a colony PCR on six different PUS381 colonies from the plates Benj prepared on 8/9.

* 10/9/19 - Wet lab

500mL cultures of PsiM and PsiK in BL21 cells were incubated, shaking, at 37C until they reached an OD~0.5. They were then induced with IPTG and moved to the room temperature shaker overnight.

Benj miniprepped the 5mL Vivid cultures he prepared the day before. He then sent the DNA off for sequencing.

Nathan prepared the PsiD colony PCR products for sequencing.

* 11/9/19 - Wet lab

Merrie streaked out colony 5 from Isobel’s colony PCR as it was the only colony to not give a band with the PUS382 primer.

Fahad pelleted the induced cultures of PsiM and PsiK from 10/9. The pellets were then stored in the cool room overnight. Fahad also made up some LB dry mix for the lab.

Benj miniprepped a 5mL wild type Vivid culture.

* 11/9/19 - Human Practices

Emma and Nathan conducted an interview with Dr Prashanth Puspanathan. Read our ‘Psilocybin Talks’ [here!](https://2019.igem.org/Team%3ASydney_Australia/Human_Practices)

* 11/9/19 - Team meeting

A bit of a difficulties troubleshooting meeting! Nathan revealed that it looks like PsiD has three deletions resulting in a nonsense mutation, made plans to do more protein ID work using larger cultures he made the day before. Isobel discussed that PUS381 has potential contamination so we need to take the colony PCR positive colony and use that to rebuild our plasmid stocks. Benj realised that the Vivid proteins on Xgal are still able to cleave the Xgal, making a product that will interfere with the fluorescence assay, will begin to replate out onto normal LB-Km plates.

However, great progress has been made on the website by Benj. Our drop down menus are coming along nicely and he aims to finish the HP page by the end of the week.

Discussed the HP interview progress - an interview with an MP is planned for later in the month, we need to interview some scalability and legal people.

Discussed who will be talking at the upcoming outreach presentations - Nathan and Emma at JAMS, Fahad and Isobel at the UNSW Symposium.

* 12/9/19 - Wet lab

Isobel decided to redo the restriction digests of the PUS381 stocks to see if any were better than those currently used to make colonies. No good results were found.

Nathan performed protein extraction on the pellets Fahad prepared on 11/9. The proteins were then run on an SDS-PAGE gel. However, it appeared that cell extract was still present in the elution columns which means that the samples require further purification.

* 13/9/19 - Wet lab

Nathan prepared two 50mL cultures each of PsiK and PsiM in BL21. Again, the cultures were shaken at 37C until they reached an OD of ~0.5. The cultures were then induced with IPTG and returned to the 37C shaker overnight.

Using the sequencing results generated from 10/9 colony PCR products, Nathan selected two colonies that did not have any deletions inside the PsiD gene to grow up in 50mL cultures for plasmid prep, and those plasmids used for BL21 transformation.

* 14/9/19-Wet lab

Nathan attempted an Ni+ column extraction of half of the cultures prepared the day before. Several different fractions were collected for running on an SDS-PAGE gel. The other two cultures were pelleted and left in the cool room.

Benj prepared another indole assay, incubating all cultures until they reached OD 0.5, adding Ala to all cultures, and inducing one culture per plasmid type (PCW, PUS381, PUS382). They were incubated at 37C overnight.

Benj also prepared one 50mL culture of PUS381 from 11/9 to use for DNA extraction to be grown overnight.

* 15/9/19 - Wet lab

Isobel miniprepped the PUS381 colony incubated the day before and performed a restriction digest on that DNA. Still no good looking results.

Isobel then reviewed the results of the indole assay and found that all cultures had turned red/brown.
This is when we started to suspect that it was the Ala that was causing the colour change, not the IPTG.

Isobel also miniprepped the positive PsiD colonies from Nathan’s colony PCR on 9/9.

Week 18: 16/9/19 - 22/9/19

* 16/9/19 - Wet lab

Isobel picked a different freshly transformed colony of PUS381 to grow up, miniprep, and restriction digest. She also ran PCW as a control as she was beginning to suspect the enzymes had been contaminated. The results continued to look suspect, so a restriction digest with different stocks of the same enzymes was planned for 18/9.

* 17/9/19 - Wet lab

Benj perfomed PCR on two CH1 Vivid variants and one wild type variant. All Vivid variants remaining on Km-Xgal plates were restreaked onto Km only plates.

Merrie performed a heat shock transformation of PK18 into Top 10 cells and plated them onto LB-Kan plates. She also prepared 5mL cultures of each of the Vivid variants and one culture of Top 10 with PK18. These were shaken at 37C until they reached an OD of 0.5. Each culture was then diluted to an OD of 0.02 and transferred to a 96 well plate to prepare for a fluorescence assay.

Merrie prepared the fluorescence and particle standard curves for the assay using an adapted version of the iGEM fluorescence protocol using the TECAN plate reader.

PsiD and PUS382 PCR products were sent off for sequencing.

* 18/9/19 - Wet lab

Fahad performed a comparative restriction digest using the stocks Isobel had used for her previous restriction digests on PUS381 and some fresh enzyme stocks. The results clearly showed that those Bam and Xba stocks had been contaminated with HindIII. Those stocks were disposed of and future experiments would be carried out using fresh, uncontaminated stocks.

Nathan transformed BL21 cells with Psi D, K, and M.

* 19/9/19 - Wet lab

Nathan checked the DNA concentrations of the plasmid stocks and reattempted the transformations done the day before. However, the transformations failed due to bad LB from an autoclaving issue.

Merrie began to prepare Golden Gate Constructs of PsiD/K/M in PUS250.

* 19/9/19 - Team meeting

Discussed the progress with the fluorescence assay, Golden gate cloning, plans for a collaboration with Macquarie, the conference banner, and Integrated Human Practices.

* 20/9/19 - Wet lab

Merrie digested, ligated, and transformed Nick’s Vivid mutants into pk18, and then into TOP10 cells to prepare for a fluorescence assay.

Nathan retransformed TOP10 cells with Psi D, K, and M. Also performed a PCR across the insert region for each of the Pet28/Psi Constructs.

Merrie also transformed TOP10 cells with the golden gate products prepared the day before.

Nathan and Isobel also began some protein work to investigate the solubility of PsiM. Initial SDS PAGE gels showed no differences in banding between the induced and uninduced samples for both the insoluble and soluble lanes.

* 21/9/19 - Wet lab

Benj patch plated 20 clearly fluorescent Vivid mutants from the transformation performed 20/9/19.

Nathan discovered that all his transformants from yesterday grew.

Week 19: 23/9/19 - 29/9/19

* 23/9/19 - Wet lab

Isobel performed colony PCR on 30 white golden gate colonies. One purple colony was picked also as a negative control.

Nathan ran some PsiK and PsiM PCR products on a gel to observe PCR results.

Isobel performed gel purification on PsiM and PsiK DNA from Nathan’s gel.

Isobel also then transformed PsiM, K, and D into new BL21 cells containing pGro7 (a plasmid coding for a protein folding chaperone).

Isobel made more golden gate transformations of TOP10 cells as no positive colony PCR results were found.

Isobel performed gel purification on PsiM and PsiK DNA to prepare for sequencing.

Merrie grew up cultures of the Vivid clones in 5mL LB broth to prepare for the fluorescence assay. After all colonies reached an OD600 of ~1, she diluted the cells to an OD of 0.02, as per the iGEM measurement protocol. Samples were loaded into a 96-well plate with eight replicates of each culture, including one Pet28-cell control, and an LB only control.

* 23/9/19 - Human Practices

Emma and Isobel conducted an interview with Alan Davis at John Hopkins. Read our ‘Psilocybin Talks’ [here!](https://2019.igem.org/Team%3ASydney_Australia/Human_Practices)

* 24/9/19 - Wet lab

Merrie cloned PsiD/K/M into PUS250 using golden gate cloning again.

* 24/9/19 - Outreach

Presented at JAMS Sydney - an amazing opportunity to showcase our project to other microbiologists and microbiology fans! Read more about the evening [here!](https://2019.igem.org/Team%3ASydney_Australia/Public_Engagement)

* 25/9/19 - Wet lab

Isobel performed colony PCR on 30 golden gate colonies from 23/9. No positive colonies were found.

* 25/9/19 - Human Practices

Emma and Isobel conducted an interview with Margaret Ross at St Vincents. Read our ‘Psilocybin Talks’ [here!](https://2019.igem.org/Team%3ASydney_Australia/Human_Practices)

* 25/9/19 - Team meeting
We discussed the Golden gate progress, beginning the new indole assay, optimised PsiH cloning, the Maquarie collaboration, and how to handle the VVD fluorescence data. On the dry lab side, we discussed the progress on the wiki, the upcoming Human Practices interviews, upcoming outreach presentations, and the team video. We also were reminded that we need to apply for special consideration for our mid-Jamboree assessments - it's all getting so close!* 26/9/19 - Wet lab

Merrie performed another colony PCR on the golden gate clones using 21 colonies. Emma patch plated the colonies ready for isolation.

Merrie also transformed TOP10 cells with the golden gate product from 24/9.

Nathan grew up cultures of PsiD/K/M to try to purify out the enzymes. Cultures were induced ~OD600 0.5. Fahad pelletted the cells and saved the pellets for protein purification.

Fahad also transformed DH5-alpha cells with pGro. Also tried to transform some DH5-alpha cells with pGro and the following: PCW, PUS381, PUS382.

Benj set up 5mL cultures of Vivid WT, CH2, CH3, CH4, and three of Nick’s error prone PCR mutants to prepare for a fluorescence assay excitation and emission spectra. Assay was performed as per the iGEM measurement protocol.

* 26/9/19 - Human Practices

Conducted an interview with Sam Bannister from the Lambert Initiative. Read our ‘Psilocybin Talks’ [here!](https://2019.igem.org/Team%3ASydney_Australia/Human_Practices)

* 26/9/19 - Outreach and Collaboration

Presented at the UNSW symposium. Helping UNSW with their outreach by educating UNSW students to both the wonder of Synthetic Biology and the sustainability considerations was an awesome opportunity.

* 27/9/19 - Wet lab

Fahad performed bead-beater protein purification on the PsiD/K/M samples and prepared them for SDS-PAGE.

Benj miniprepped the Vivid WT, CH2, CH3, CH4, and error prone PCR mutants that were used in the fluorescence assay.

* 28/9/19 - Wet lab

Nathan ran Psi D, K, and M protein extracts on an SDS-PAGE gel after separating based on solubility. Protein was observed, but only in the insoluble fractions.

Isobel checked the transformations of DH5-alpha cells with pGro. The pGro only transformation and the pGro/PCW transformation worked but the others did not. She decided that the pGro-transformed cells needed to be made competent again. She began step one of that process.

She inoculated 3mL of the 5mL overnight cultures of the DH5-alpha-pGRO cells and top10 cells into 100mL LB and performed the chemical competency protocol on the cells. 100uL aliquots were plated onto plates of the appropriate agar and incubated overnight.

* 29/9/19 - Wet lab

Isobel performed the transformed various plasmids for the Macquarie collab into DH5-alpha cells.

She also tested the competent cells made on 28/9 by transforming them with PCW. These plates were then incubated overnight.

Week 20: 30/9/19 - 5/10/19

* 30/9/19 - Wet lab

Fahad prepared cultures of PsiD/K/M for induction. Cultures were induced when they reached an OD600 of ~0.5.

Merrie miniprepped cultures of the two positive golden gate colonies. The DNA was stored in the cold room for PCR the next day.

Merrie also prepared overnight liquid cultures of all the Macquarie collaboration constructs.

* 1/9/19 - Wet lab

Fahad decanted the overnight cultures of PSID/K/M (+/-) into 15ml falcon tubes and centrifuged 3900 for 15 min to pellet. The cells were resuspended in 5ml of wash buffer A by vortexing and were again pelleted by centrifugation 3900 for 15 min.

Nathan set up PsiD/K/M cultures at 37C shaking until they reached an OD600 of ~0.5. The cultures were then induced and incubated at 37C, 30C, and room temp after induction. After five hours all cells were pelleted, and the pellets were stored in the cold room.

Merrie performed spanning PCR on the golden gate constructs from 30/9. Both colonies were positive.

Merrie moved the Macquarie collaboration cultures to the cold room, prepared 0.1mM IPTG for induction, and prepared the LB-antibiotic mastermixes. The samples were put into the TECAN for the fluorescence assay, but it was aborted after ten hours as the cultures maxed out the machine.

* 2/10/19 - Wet lab

Merrie did PCR on codon-optimised PsiH to add the restriction sites to move it into PUS381.

Nathan resuspended the cell pellets from 1/10 in Buffer A, then centrifuged, then resuspended in TE. Soluble extracts were obtained by beadbeating and nanodropped. Cell extracts were then stored in the cold room overnight for SDS-PAGE.

Merrie also ran a gel on the PCR products from 1/9 to perform gel purification.

* 2/10/19 - Team meeting

We discussed the attributions page, collaborations page, and the Human Practices progress. We feel as though we have these aspects of the progress very much under control. We discussed where to characterise our WT-VVD and VVD-Met130 mutant parts in terms of the medal criteria. We also decided on a lab work cut off, October 16 for most wet lab work, October 23rd for characterisation wet lab work for our presentation and poster.

* 3/10/19 - Wet lab

Isobel digested codon-optimised PsiH (CO-PsiH) and PUS381 with Bam and Xba. Isobel then ligated CO-PsiH and PUS381. This mixture was then transformed into DH5-alpha/pGRO cells.

Nathan heated the SDS page gel of protein extracts from 2/10/19 3 minutes at 95°C before loading onto SDS page gel 10μg of protein per lane plus SDS buffer and water to make up 20μL.

Merrie performed gel purification on the gel ran yesterday. The concentration was not high enough after purification so another PCR and gel purification was performed. This second gel purification resulted in yields suitable for sequencing. The samples were set up and sent off for sequencing.

* 4/10/19 - Wet lab

Isobel performed colony PCR on 28 CO-PsiH-DH5alpha/pGRO colonies.
5 positive colonies were found. Decided to call these codon-optimised PsiH/PUS381 constructs PUS383 from now on.

Nathan ran 15°C soluble cell extracts on SDS page again heated for 3 minutes at 95°C before loading onto SDS page gel 10μg of protein per lane plus SDS buffer and water to make up 20μL.

Emma performed heat shock transformation of the golden gate constructs (14 and 16) in PUS250 - henceforth referred to as PUS387-14 and PUS387-16 - into DH5-alpha-pGRO as per the lab protocol. PUS250 was also transformed as a positive control. The transformations were plated on LB-Km-Cm plates and incubated overnight.

Merrie started new overnight cultures of the Macquarie constructs for a new fluorescence assay.

* 4/10/19 - Human Practices

Fahad met with Adamn Bandt, the Federal Greens spokesman for science, in Melbourne to discuss the responsible and democratic use of GM technology, especially in reference to our project. This chat was part of our Psilocybin talks series on our [Human Practices page!](https://2019.igem.org/Team%3ASydney_Australia/Human_Practices)

* 5/10/19-Wet lab

Isobel began making the indole assay pre-induction media. Inoculated 3mL bottles of the media with each of PUS381, PCW, PUS382, and three different PUS383 colonies that had positive PCR results the day before.

Isobel also performed a spanning PCR on the PUS383 colonies to determine which colony will be used for the indole assay. All three colonies had an insert of the correct size.

Merrie spun down the 30°C and 37°C induced and unindexed samples from PSIK and PSIM and replaced the TE with SDS loading buffer. They were then transferred to bead beating tubes and beat for 30s, iced for 2 min, and spun at max speed for two minutes. They were then transferred to new tubes on ice.

Nathan made 3x 500ml of LB Km/Cm with added arabinose to 2mg/ml. 50ml cultures of PSID, PSIK, and PSIM were used to seed the 500ml conical flasks. OD was measured periodically until it reached approx OD600 0.5. They were then induced with IPTG and returned to the 15C shaker.

Merrie made up 10mL liquid cultures of the PUS387 transformations from 4/10/19. Aliquots were taken to use to assess psilocybin production.

Merrie set up another fluorescence assay for Macquarie according to their protocol but with some minor changes. The cultures were pipetted uninduced, then the IPTG was added. She also changed the excitation to 480, the emission to 525, and the gain to 75, to better prevent maxing out the TECAN. The machine was set to perform 140 cycles, 30 min apart, shaking at 150rpm, 37 degrees celsius.

Merrie also made glycerol stocks of the error-prone PCR Vivid mutants to put in the -80C freezer.

Week 21: 6/10/19 - 13/10/19

* 6/10/19 - Wet lab

Isobel started the indole assay. To TB+additives bases, 0.5mL of each pre-induction culture was added: PCW, PUS381, PUS382, and PUS383.

Isobel also prepared a 50mL LB culture of PUS383 to prepare for a mini-prep.

Fahad prepared a 400mg/mL arabinose stock to induce the indole assay cultures. He then induced the cultures in the evening with arabinose, IPTG, and alanine.

Isobel streaked PSIDKM in Dh5a pGro7 onto kan/cm plate from bottle of culture in cold room put in 37 degree C incubator.

Fahad removed PSIK/D/M cultures from shaking incubator (15°C). The cultures were then decanted into 11x falcon tubes 50ml each and centrifuged 3900 for 15 min to pellet. The supernatant was poured off and cells resuspended in 5ml wash buffer A then decanted into single falcon tubes for each plasmid and centrifuged at 3900 for 15 minutes. Again, the supernatant was poured off and cells resuspended in 5ml 1x TE supplemented with 3ul of 10x protease inhibitor then frozen at -30C.

* 8/10/19 - Wet lab

Merrie miniprepped the PUS383 50mL LB culture.

Merrie also discovered that the PCR and sequencing of PUS387-14 and PUS387-16 failed due to the use of an incorrect primer. She decided to redo the PCR using the correct primers and a PUS250 and no template control.

Isobel checked the results of the indole assay. PCW changed colour very drastically, but no colour change was observed in any of the other cultures. One set of cultures was set aside to be used for protein purification. Protein purification was done according to the protocol for insoluble proteins, and the resulting protein extract was run on an SDS-PAGE gel. Extra protein extract was frozen in TE before SDS-loading buffer was added. The gel was not the best.

Merrie performed a heat shock transformation according to Coleman lab protocol colony 14 PUS387 into DH5a pGro7, she also prepared a NTC and PET28 positive control.

She also performed a PCR on PUS387-14 and 16, with PUS250 and a no template control.

* 9/10/19 - Wet lab

Isobel transformed DH5-alpha cells with PUS381/2/3 to prepare for a hydroxytryptamine assay. She also co-transformed DH5-alpha with PUS387(14/16) and PUS382/3.

She then also prepared PUS383 for sequencing.

Merrie took the golden gate transformation plates from 8/10 out of the incubator and used them to inoculate 2x 10mL cultures of TB for a single plasmid psilocybin assay. She realised that she had transformed the wrong control (pet28 instead of PUS250) and would have to repeat with PUS250 later. They were incubated for several hours, but overshot their required OD. They were diluted and returned to the incubator until they reached the required OD of 0.5. They were then induced with 10ul of 100mM cumate and placed in 15 degree shaker in 50ml falcons. After 7 hours incubating they were spun down and resuspended in phosphate-glucose buffer and 4-hydroxytryptamine was added. One tube of each plasmid was placed in the 37C shaker, and one in the 15C shaker.

Merrie also ran the results of yesterday’s PCR on a gel.

Nathan ran Fahad’s protein samples from 6/10 on an SDS page gel to visualise protein expression and solubility at 15C. Gels were stained with Coomassie blue.

He also performed dialysis of the his-tagged purified proteins using the cell pellets Fahad prepared according to protocol.

Isobel put Nathan’s gels in the high and low destain.

* 9/10/19 - Team meeting

Discussed the results of the indole assay, the upcoming scale-up experiment, the upcoming phosphate assays, and the need to send all our protein gels for LCMS. Fahad revealed that the Codonator would soon be up on the wiki - very exciting! Emma begged for help transcribing the recent interviews. Isobel offered to transcribe Margaret Ross's interview, Fahad his interview with Adam Bandt, and Nathan offered to transcribe the Dale interview. Merrie said that she has fully finished the work required for the Macquarie collab, she has sent them our data and we have received their gel. Benj taught us how to upload images for the wiki. We discussed the upcoming deadlines, Fahad offered to take the lead on the making and populating of the parts pages.

* 10/10/19 - Wet lab

Fahad retrieved the extra PUS381/2/3 protein extract from the freezer and performed another SDS page on the samples to get a better image.

Emma inoculated the PUS381/2/3 and PUS387-PUS382/3 cultures transformed yesterday into antibiotic-glucose-LB media. They were put into the 37C shaker overnight. Emma also prepared the TB for the next step of the hydroxytryptamine and psilocybin assays and sent it for autoclaving.

Merrie decided to repeat yesterday’s single-plasmid psilocybin assay with the correct controls and no shaking. She also spun down the tubes from the day before and found that PUS387-14 and 16 had black pigment to their pellets. The cells were stored in glycerol in the -20C freezer.

She also performed a miniprep of PUS387-14 and 16 to prepare for sequencing. The samples were prepared with primers and sent off for sequencing.

Nathan realised he left the PsiM fractionated samples out overnight (big sad). He ran all three on gels regardless.
Nathan decided with the help of our supervisor, Nick, to abandon the in-vitro enzyme assay of PsiM due to it being likely degraded and didn’t purify well. He dialysed PsiD and PsiK again at 4C overnight, stirring.

Nathan and Benj performed a scale-up exercise in collaboration with the chemical engineering department. Nathan inoculated a 1.5L TB culture with DH5-alpha PUS250 cells. He took OD600 every 20mins.

Benj did the same but using the 1.5L incubator in the engineering department and measuring every 30mins.

* 11/10/19 - Wet lab

Merrie transferred the PUS381/2/3 and PUS387-382/3 cultures from the LB-glu media into the TB and added the additives from the indole assay. All cultures were incubated at room temp until induction.

Merrie induced the PUS381/2/3 cultures after six hours with IPTG and arabinose and left them shaking at room temperature, then induced the PUS387-PUS382/3 cultures with IPTG, arabinose, and cumate and moved them to the 15C shaker.

Fahad thawed more of the PsiD cultures frozen on 6/10. 10mL aliquots of these were decanted into 50mL falcon tubes, washed in PBS 5ml, and then after the supernatant was poured off. Cells were then resuspended in 5ml fresh phosphate buffer with 1mM tryptophan. Took 1ml supernatant and froze. Remainder spun down and returned to shaker.

Nathan performed some transformations of PUS250, PUS250/PUS381, PUS381, and PUS381/Pet28-PsiM, with a no template control into DH5-alpha-pGRO cells. They were plated and put in the incubator overnight.

In the evening (six hours after induction), Isobel pelleted the cells from the PUS381/2/3 assays and the PUS387-PUS382/3 assays and resuspended them in phosphate-glucose buffer. All cultures had their required intermediates added. Isobel also took time point zero samples. These were stored in the freezer to prepare for LCMS.

* 12/9/19 - Wet lab

Isobel performed a PCR to check 381/382/383 samples are correct due to suspicious OD600 results yesterday. The results showed that the cultures were in fact correct.

Merrie performed a restriction digest according to lab protocol of PCW, and PUS381/2/3 with Bam & XbaI to get a nice gel image for the wiki. However the gel showed incomplete digestion so would have to be repeated.

Nathan removed the dialysis tubes from the fridge and extracted 1mL of PsiD and PsiK into separate 1mL eppendorfs to prepare for the in vitro assay. In one tube there was 200uL enzyme alone, in another tube there was 200uL enzyme with the substrate. For PsiK, 5mM ATP MgCl2 was added to both tubes. Reactions were left to proceed for two hours before being moved to the freezer to wait for LCMS.

Emma span down the PUS381/2/3 and PUS387-PUS382/3 cells and took 1mL samples of the supernatant as our t=24 samples, these were stored in the freezer to send off for LCMS.

* 13/10/19 - Wet lab

Merrie performed another restriction digest on PCW and PUS381/2/3 according to protocol, but was still having troubles with incomplete digestion.

Merrie removed the PUS387 single plasmid psilocybin assays from the 15C shaker. The supernatant was removed and placed in the freezer until it could be sent off for LCMS.

* 13/10/19 - Outreach

Benj, Emma, Isobel, and Nathan set off for Brisbane to participate in the Synthetic Biology Australia (SBA) conference!

Week 22: 14/10/19 - 20/10/19

* 14/10/19 - Wet lab

Merrie prepared the protein samples for PsiD/K/M for LCMS. She also sent our enzyme assay supernatants off for LCMS.

Merrie also began work on the bSFP by reconstituting the G-block to clone it into pk18. The ligation mixture was transformed into TOP10 cells and plated onto LB-Km-Xgal plates.

* 14/10/19 - Outreach

Day one of the SBA conference! Nathan, Isobel, Benj, Emma, and Fahad attended.

* 15/10/19 - Wet lab

Merrie made 5ml cultures of bsfp, WT vivid, CH1, CH2, CH3, Met130-10 and put them in the 37 shaker overnight to prepare for the characterisation fluorescence assay. She was concerned by the small number of white colonies observed on the bSFP plates so she planned another transformation as a back up.

* 15/10/19 - Outreach

Day two of the SBA conference! Nathan, Isobel, Benj, Emma, and Fahad attended.

* 16/10/19 - Wet lab

Merrie performed another transformation of bSFP into TOP10 cells. She also set up the fluorescence assay with the overnight cultures from the day before as per the iGEM measurement protocol.

Merrie also performed a digest of PCW，PUS381/2/3 and ran the products on a gel to make a nice gel for the Wiki. However the gel was not so nice, so it needed to be performed again.

* 16/10/19 - Outreach

Day three of the SBA conference! Emma and Isobel presented our progress so far. The presentation went fantastically and they fielded questions well.

* 18/10/19 - Wet lab

Merrie received the results from the bSFP fluorescence assay. The levels were surprisingly low so she decided to redo the digestion and ligation of the G-block into pk18. The new ligation mix was then transformed into TOP10 cells and plated on LB-Km-Xgal plates.

* 19/10/19 - Wet lab

Isobel and Merrie performed colony PCR on bsfp using NVC15b and 16b plus patch plate and 5ml cultures of 16 white colonies.

They also set up 5ml cultures of CH1 and WT Vivid, PK18 and Met-130. Then they did another transformation of TOP10 cells with the ligation product from yesterday.

Nathan digested each pET28-psi construct using HindIII and EcoRI to make some nice gels for the wiki.

* 20/10/10 - Wet lab

Nathan performed a double restriction digestion of pUS250 and pUS387 clones (14 and 16) using NEB's HindIII-HF and EcoRI-HF enzymes.

Merrie attempted to make more cultures of bsfGFP but they were not growing. They were abandoned as our comparative protein for our VVD\_Met130 experiment.

Week 23: 21/10/19 - Wiki Freeze!

* 21/10/19 - Wet lab

Merrie Prepared cell lysates of MET130I and sfGFP for thermal stability assay and performed assay with StepOnePlus Real time PCR system. She also measured the excitation and emission spectrum of the MET130I fluorescent protein using the Tecan Spark plate reader.

* 21/10/19 - Team meeting

The team met to finalise details on the wiki and parts registry. Results from the melt curve assays were added to the wiki and registry.

And with that, all we needed to do was polish off our poster and presentation, and get ready to head off to the Jamboree! If you want to learn more about what we did in our final days of the project, check out our social media.


[![](https://2019.igem.org/wiki/images/8/8b/HQ_page_logo.jpg)](https://2019.igem.org)

# Team:Sydney Australia/Experiments

Navigation

[![](https://static.igem.org/mediawiki/2019/c/cb/T--Sydney_Australia--MagiColi.png)](https://2019.igem.org/Team%3ASydney_Australia)

* Team

  [The Team](https://2019.igem.org/Team%3ASydney_Australia/Team)
   [Collaboration](https://2019.igem.org/Team%3ASydney_Australia/Collaborations)
* Project

  [Description](https://2019.igem.org/Team%3ASydney_Australia/Description)
   [Design](https://2019.igem.org/Team%3ASydney_Australia/Design)
   [Experiments](https://2019.igem.org/Team%3ASydney_Australia/Experiments)
   [Notebook](https://2019.igem.org/Team%3ASydney_Australia/Notebook)
   [Characterisation](https://2019.igem.org/Team%3ASydney_Australia/Contribution)
   [Results](https://2019.igem.org/Team%3ASydney_Australia/Results)
   [Demonstrate](https://2019.igem.org/Team%3ASydney_Australia/Demonstrate)
   [Improve](https://2019.igem.org/Team%3ASydney_Australia/Improve)
   [Attributions](https://2019.igem.org/Team%3ASydney_Australia/Attributions)
* Parts

  [Parts Overview](https://2019.igem.org/Team%3ASydney_Australia/Parts)
   [Basic Parts](https://2019.igem.org/Team%3ASydney_Australia/Basic_Part)
   [Part Collection](https://2019.igem.org/Team%3ASydney_Australia/Part_Collection)
* [Safety](https://2019.igem.org/Team%3ASydney_Australia/Safety)
* Human Practices

  [Human Practices](https://2019.igem.org/Team%3ASydney_Australia/Human_Practices)
   [Outreach & Engagement](https://2019.igem.org/Team%3ASydney_Australia/Public_Engagement)
* Achievements

  [Medals](https://2019.igem.org/Team%3ASydney_Australia/Medals)
  [Modelling](https://2019.igem.org/Team%3ASydney_Australia/Model)
  [Software](https://2019.igem.org/Team%3ASydney_Australia/Software)

[![](https://static.igem.org/mediawiki/2019/c/cb/T--Sydney_Australia--MagiColi.png)](href%3D)

[![](https://static.igem.org/mediawiki/2016/9/96/T--Sydney_Australia--newusydlogo.png)](https://sydney.edu.au/science/schools/school-of-life-and-environmental-sciences.html)

![](https://static.igem.org/mediawiki/2019/f/f9/T--Sydney_Australia--Sponsors2.png)

Experiments

# Key Experiments

Characterisation of PsiD/K/M in Vitro

#### Aim

To confirm the production and activity of the PSIK, PSID and PSIM in individual constructs prior to combination in single inducible vector.

![](https://static.igem.org/mediawiki/2019/3/3a/T--Sydney_Australia--PSIproteincharacterisation.png)

Assembling PsiD, K, M Gene Cluster for Magi.Coli

#### Aim

To confirm the production and activity of the PSIK, PSID and PSIM in individual constructs prior to combination in single inducible vector.

![](https://static.igem.org/mediawiki/2019/c/c5/T--Sydney_Australia--PsiDKMGeneCluster.png)

Expressing and Characterising PsiH Enzymes

#### Aim

This part of the project aimed to express and characterise the activity of the PsiH, a P450 mono-oxygenase enzyme that converts tryptamine to 4 hydroxytryptamine in vivo before putting it into our single-cell system.

![](https://static.igem.org/mediawiki/2019/8/87/T--Sydney_Australia--PsiHStuff.png)

Creation of Two-Plasmid System

#### Aim

To transform our two plasmids into the same cell so the entire pathway from the mushroom can be recreated functionally in E.coli.

![](https://static.igem.org/mediawiki/2019/2/24/T--Sydney_Australia--filename_twoplasmidassay.png)

Culturing the Magi.coli Chassis: Lab Vs Fermenter

#### Aim

![](https://static.igem.org/mediawiki/2019/3/3c/T--Sydney_Australia--CulturingChassis.png)

Codon Harmonisation Experiments with Vivid

#### Aim

The aim of this experiment was to determine which type of codon harmonisation/optimisation (rank-order method, absolute frequency method, relative frequency method and IDT codon optimisation) would improve the expression of the VVD protein compared to the wild-type codon usage. The results of this experiment would then be used to codon harmonise our psi genes to improve their expression.

![](https://static.igem.org/mediawiki/2019/f/f6/T--Sydney_Australia--VividThings.png)

Error-Prone PCR of Vivid

#### Aim

To increase the fluorescence of the IDT-codon optimised VVD to further increase its utility as an iLOV-based blue fluorescent protein.

![](https://static.igem.org/mediawiki/2019/9/93/T--Sydney_Australia--ErrorPronePCRVivid2.png)

# Protocols

General Lab Safety and Aseptic Technique

#### Innoculating or Transferring Cultures

* Wipe down the bench with ethanol or F10FC before starting.
* Work close to the Bunsen burner.

+ Use the hot ‘blue flame’ to get a strong updraft, flame the necks of bottles, and
  to flame your loop (if using a metal loop, do not flame plastic loops, you will
  have a bad time).
+ The safety flame (yellow) is only good for seeing when you have lit your
  bunsen burner and for lighting the ethanol on your glass spreader, if using.
  Again, do not light plastic spreaders on fire - you will have a bad time.

* Keep the work area clear of clutter, ensure everything you need is within easy
  reach such that no part of your body will need to cross the flame. You do not want
  to set on fire either.
* Don’t do culture work on top of your lab book or other paper notes. These should
  be kept in green spaces. Paper near aseptic work increases the risk of fire, and
  your beautiful notes going into the autoclave.
* Flame the neck of glass bottles and tubes when opening and closing them

+ Pass through flame back and forth briefly three or four times over three seconds.

* Be mindful of where you put things, act as though the bench is lava.

+ i.e. once you remove a sterile plastic loop from the packet, or flame your metal loop, do not put it back down on the counter or it will have to be re-flamed or disposed of.
+ Do not contaminate your workspace by putting loops with culture on them

* If your lab coat sleeves hang from your wrists (i.e. are not elasticated so they fit
  snugly on your arm), do not allow them to flap around. Tuck them into your gloves
  if possible, tape them or elastic band them to fit snugly, or roll them up if not
  working with corrosive or toxic reagents.

+ Fun fact: loose sleeves + bacteria + flame = contamination or fire.

##### Sterile Bottles, Solutions, and Gear

* Label EVERYTHING carefully! Write chemicals used, plasmid names, media type, date, and your name.
* Make sure any hazardous reagents are clearly labelled with the chemical name, the
  concentration, and the nature of the hazard (e.g. flammable/corrosive/toxic). Store
  all hazardous reagents in the appropriate area.
  DO NOT leave unlabelled hazardous chemicals on the bench. You never know if
  any of your labmates are feeling thirsty and decide to ignore the ‘no drinking
  rule’. Do you want their dumb deaths on your hands?
* When you take lids off, do not put them down on the bench unless
  necessary. If you need to put the lid down (weakling), put them upside down, so the
  screw thread is facing upwards. Surfaces are a huge source of bacterial
  contamination.
* Don’t leave lids of bottles for any longer than necessary. Turns out the air is lava
  too.
* Getting sterile Eppendorf tubes out of the autoclave beaker:

+ OPTION 1: take off foil top, carefully pluck out the tube by its base or edge, place
  carefully in a tube rack, repeat if necessary, replace foil.
+ OPTION 2: take off foil top, shake out tubes into the foil, carefully place in the tube
  rack - being careful not to touch the inside of the tube, replace foil.
+ DO NOT : rip off the foil, plunge your hand into the beaker, rummage around
  like it’s a bargain bin, emerge triumphantly with a tube, then leave the foil in
  disarray until some poor sad soul realises they need to send the whole batch
  down for autoclaving again because of your inconsiderate behaviour.

#### Pipetting with Gibsons

* Wipe down your pipettes with ethanol every. single. day. If you do not do this, I
  really don’t think you can call yourself a Molecular biologist! (luckily for me I identify as a microbiologist so I’m safe)

+ Also do this when changing from microbial to DNA work, or if you’re concerned
  you (or someone you know) have contaminated your pipette somehow.

* If you’re having persistent issues with contamination, do not just try to dump your
  pipette on someone else’s bench and take theirs. There is another way!

+ Dissemble the pipette according to the manufacturer’s instructions.
+ Soak in bleach (0.5% hypochlorite = a 1/10 dilution of household bleach, freshly
  prepared), so that all parts of the pipette are covered.
+ Leave to soak for 30min. Do not leave for longer or you are at risk of the pipette parts rusting.
+ Rinse extensively with water, then with 80% ethanol.
+ Allow to dry.

* Remember that just the tip is sterile! No matter how recently you have swabbed
  your pipette with alcohol, the main body of the pipette is still contaminated in some
  way (DNA, microbes, enzymes, they’re everywhere! Remember - the air is lava).

+ The body of the pipette should not come into contact with anything that needs
  to remain sterile (e.g. the inner walls of bottles or falcon tubes).

* Hold the pipette vertical! Or as close to it as you can get. If you hold it sideways the
  liquids may run up the barrel of the pipette, damaging the inner workings, or worse
  - contaminating your culture or PCR with whatever gunk got up there before!
* As long as you are careful with your pipetting, most work will not require plugged
  pipette tips.

+ They are only really required for RNA work and 16S RNA PCRs as these
  procedures are incredibly sensitive to DNA/RNA contamination.

* Be careful when getting tips out of the box, we don’t want any tip casualties. Open
  the box the whole way, put the tip on firmly, move the pipette away, then close the
  box.
* Don’t hit the tip on the lid of the box, don’t leave the box open long after you’ve
  got the tip out, and don’t go around knocking other tips on your way out like
  some sort of pipetting Godzilla!
* All the pipette tip wants is to be part of an experiment, don’t let them die
  before they get their time in the spotlight.

#### Setting up PCRs and Other Enzyme Reactions

* Preparation prevents poor performance! Swab down your bench and pipettes with
  ethanol, label your tubes, and most importantly - read the protocol carefully!
* Keep all enzymes in the freezer or on ice at all times. No exceptions! (except maybe like three
  exceptions) Restriction enzymes, ligase, polymerases, reverse transcriptase, phosphotase,
  kinase, Gibson assembly mix, etc.
* Keep all DNAs (plasmids, PCR products, genomic DNAs) either in the fridge or
  freezer when they are not actively being used. DNA can sit on the bench for a little
  while (few minutes up to an hour), but don’t abandon it in the harsh outdoors.
* Double check all buffer and incubation requirements. It may help to write things
  such as times and temperatures on a separate piece of paper as you read through
  the protocol. That way you can make a list and check it twice!
* Make up the bufer first upto to 1x concentration or what is needed for your experiment, THEN add your enzymes and/or DNA.  Least expensive to most expensive as I say \*wink\*.
* Handle your tubes carefully.

+ Don’t touch any part of the inside of your tubes, including the inside part of the
  lids with your gloves or gross human fingers. Eww.
+ Be careful when opening the tubes especially as your thumb can easily slip
  onto the inside of the lid without realising! Try using your thumbnail, not your
  whole thumb, to carefully flick open the lid.
+ Be extra careful when putting the tubes on ice - the ice must not get inside
  your tubes! Don’t push the tubes too far down into the ice, handle them
  carefully. It can be helpful to use a ‘floaty’ (thin foam tube holder) to keep your
  tubes from going too far into the ice.

Making Antibiotic Stocks

When doing molecular biology experiments using bacteria, the ability to properly
make antibiotic stocks is life and death. Too weak, and you risk mutating
untransformed cells into antibiotic-resistant mutants that could accidentally be
unleashed on the world and kill us all… too strong… and you ruin your experiment by
killing everything and have a really bad day.

To make your time and effort worthwhile, try to make 50 or 100mL of stock at a time
to be frozen in 1mL aliquots. These steps will reference quantities for a 50mL batch,
but this can be scaled up or down depending on your available antibiotic-stock-making equipment and your antibiotic needs.

#### Typical Antibiotic Concentrations: Stock & Usage

* Ampicillin: 100 µg/ml in broth —> 100mg/mL in stock
* Chloramphenicol: 25 µg/ml (usually - 12.5 µg/ml for fosmids) —> 25mg/mL in stock
* Kanamycin: 50 µg/ml —> 50mg/mL in stock

- Tetracycline: 10 µg/ml (light-sensitive! wrap plates/broths/stocks in foil) —> 10mg/mL in stock (Foil wrapped or in black-opaque tubes)
- Streptomycin: 200 µg/ml —> 200mg/mL in stock
- Gentamicin: 10 µg/ml —> 10mg/mL in stock

#### Protocol

1. Grab your supplies. For a 50mL batch you will need:

* One falcon (50mL) full of ultra-pure water (i.e. MilliQ)
* Two empty falcon tubes (50mL)
* One 50mL syringe - plastic tip, not metal as we don’t want needle stick injuries
* One filter steriliser tip to fit your 50mL syringe
* Your antibiotic powder of choice
* A teeny tiny spoon
* A falcon tube holding rack

2. Place your falcon tube holding rack and one empty falcon on your scales, set
   scale to zero.
3. Weigh your dried antibiotic to make up desired concentration (e.g. 250mg
   Kanamycin, 5000mg Ampicillin), using the all-important teeny tiny spoon to
   measure into the empty falcon.
4. Using a measuring cylinder, measure out 50mL of your ultra-pure water. Pour
   measured water into the falcon with the antibiotic powder.
5. Seal the falcon tube and shake it like a polaroid picture until the powder is
   dissolved. Top tip: make sure that the tube is sealed or you will get antibiotic water
   everywhere!
6. Now we need to filter sterilise the stock, so prepare your other empty falcon tube
   by placing it into the rack. Open your filter steriliser tip, affix it to the end of your
   syringe, remove the plunger so you can pour your stock into the syringe.
7. Don’t leave the plunger on the bench, keep it in your hand - remember, the bench
   is lava!
8. Once the syringe is full, you can put the plunger back into the syringe and push
   your stock through your filter steriliser.
9. You have now made a stock! Congratulations! It’s time to make your aliquots!
10. Get out 1.5 mL tubes and label them with the antibiotic name/abbreviation.
11. Pipette 1mL of antibiotic into each tube. You may end up not having enough antibiotic to put into all 50 of the tubes,
    sometimes you lose an mL or so along the way. This is normal. Take deep breaths
    and just be proud you managed to make at least 45 tubes.
12. Place into a labelled ziplock bag and freeze at -20C until required.
13. Stocks can be defrosted by warming in hands as required.

Making Agar Plates

If you're going to do any Synbio, you're bound to need a TONNE of plates. Here's the best method we've found for making them!

1. Figure out the total number of plates you will need, then multiply x 25 to get the approx. amount of agar you will need (in ml). It's better to make more than you think you need! Alternatively, just make up the agar in multiples of 400 ml, this a standard amount that will make ~16 plates (this is about right for one ‘sleeve’ of sterile Petri dishes, which have 20 dishes in them).
2. Add all the required media ingredients (eg. for LB agar, this is 10 g tryptone, 5 g yeast extract and 5 g of NaCl per litre) – make sure you correct these ingredient weights for the amount of agar you are making (e.g. multiply by 0.4 if you are only making 400 ml).

DON’T ADD THE AGAR YET! (this doesn’t dissolve, it settles out)

DON’T ADD ANTIBIOTICS, TRACE METALS, or TWEEN yet, if these ingredients are needed.

3. Dispense the media into multiple lots of 400 ml in 500 ml media bottles. Make sure that the media bottles have a plastic pouring ‘lip’ on them, not just a naked glass top; the lip is important to be able to pour the agar neatly. You could also make multiples of 200 ml in 250 ml bottles, but don’t make >400 ml in a 1 L bottle – this is too heavy to easily pour plates with.
4. Once you have dispensed the media into the bottles, now you can add the appropriate amount of agar to each bottle. This is 17 grams per litre (=6.8 grams per 400 ml).
5. Add bottle caps. Don’t screw these on all the way, leave them a bit loose. Hold the caps in place with a small piece of autoclave tape.
6. Autoclave the agar. If the agar media will not be autoclaved on the same day, keep it in the cold room until the next day (microbes will start to grow within a few hours, especially in rich media like LB).
7. When the agar comes out of the autoclave, if it is still molten and very hot, put it in the hot water bath (~60°C) until it cools down to pouring temperature (~15 min). If you don’t want to use it immediately, leave on the bench until it solidifies (if it hasn’t already), then tighten the cap and it can be stored indefinitely at room temp. If agar has solidified and you want to use it straight away, microwave it (lid loose!) for approx 10 min on 50% power (these settings are for 400 ml of agar at room temp, adjust accordingly!), then put in hot water bath for ~15 min.
8. Prepare laminar flow hood for plate pouring as follows: give it a blast of UV for ~15 min then turn off UV light,), take out the doors, then turn on the light and fan. Swab down with 80% ethanol or F10SC.
9. If you don’t have access to the laminar flow hood, you can make plates on your regular lab bench. This is usually fine, but swab down the bench with ethanol first, and work close to the Bunsen flame. (if you swab your gloves with ethanol, be careful to let this evaporate before turning on the Bunsen burner!)
10. Collect one bottle of molten agar from the waterbath, wipe down the outside with paper towel, and take to your bench. Add any “after-autoclaving” additions at this stage, from filter-sterilised stock solutions (this may be antibiotics, or trace metals solution, or Tween etc). Mix by swirling for 10-20 seconds, don’t shake the bottle as this will cause persistent bubbles. Its good practice to write all the additions on the bottle, then tick them off after each addition. Forgetting these is bad!
11. Take the final agar to the laminar flow hood. Open up one sleeve of plates (at the ‘base’ end), and slide them out. Keep the sleeve! Label all the plates with the type of medium (e.g LB-Cm25). Pour the agar into the first plate; use a single smooth pouring motion. Stop when it reaches all the edges. Put the lid on. Do the next plate. Repeat. Don’t stack up the plates in big towers, they will take longer to cool.
12. Leave the plates for about 10 min to allow the agar to solidify. Then invert the plates with the lids off to allow them to dry for 20-30 min.

If you don’t dry the plates, you won’t get nice spread-plates, and the risk of contamination is greater from other microbes ‘climbing’ into the plate over the edge.

13. Pack the plates back into the sleeve, then label the sleeve with the type of agar and the date of manufacture and your name.
14. Store plates in the cold room. They should stay good for >6 months if you have been careful!

Plasmid Mini-prep Protocol for 5ml and 50ml Cultures

This protocol is based on the Coleman Lab protocol.

The solutions needed for the plasmid prep are listed at the end of the protocol. These should work with any kind of silica-based purification column and could also be adapted to purification using free silica particles (e.g. diatomaceous earth).

Wear appropriate PE including lab coat gloves and safety glasses. Hazards include NaOH, SDS and guanidine.

#### Method for 5mL Culture

1. Inoculate 5 ml of LB broth containing the appropriate antibiotic with a loopful of E.coli culture from a fresh plate of the same medium (‘fresh’ here means less than ~2 weeks old). Grow overnight (16-24 h) at 37°C with shaking.

It is critical to remember to add the antibiotics to the LB broth. Some common antibiotics and their typical concentrations are shown below:

* Ampicillin: 100 μg/ml
* Kanamycin: 50 μg/ml
* Streptomycin: 200 μg/ml
* Chloramphenicol: 25 μg/ml (usually) or 12.5 μg/ml (for fosmids)
* Tetracycline: 10 μg/ml (light-sensitive! wrap plates/broths in foil)
* Gentamicin: 10 μg/ml

2. Centrifuge the culture at 3000 rpm for 10 minutes.
3. Pour off supernatant into culture waste and resuspend the cells in 1 ml of buffer EB by vortexing. Transfer to 1.5 ml Eppendorf tube. Centrifuge 1 min at ~10,000 g. Pour off supernatant, keep pellet.
4. Resuspend the cell pellet in 200 μl buffer P1 by vortexing.
5. Add 300 μl buffer P2. Mix by rapidly inverting the tube 10 times. Incubate 10 min at room temp. (No longer than 10 min!) The mixture should go clear and become viscous as the cells break open.
6. Add 400 μl buffer N3. Mix by rapidly inverting the tube 10 times. It is essential that the N3 buffer is thoroughly mixed in. Viscosity should disappear, and a white precipitate should appear. Incubate for 5 minutes at room temp. Centrifuge for 5 min at ~10,000 g.
7. Place a spin column (silica-based e.g. ‘Econospin’ brand) into its collection tube (if it isn’t already set up that way), and carefully pipette 750μl of the plasmid extract supernatant onto the column. Avoid the white pellet and any white junk that may be floating on top. Centrifuge 30 seconds at ~10,000 g. Discard flow-through into culture waste.
8. Add 700 μl buffer PB to the column, centrifuge 30 sec at ~10,000 x g. Discard flow-through.
9. Add 700μl buffer PE to the column, centrifuge 30 sec at ~10,000 x g. Discard flow-through.
10. Repeat step 9.
11. Spin the ‘empty’ column one more time for 2 min at ~10,000 x g to remove the last traces of PE.
12. Discard collection tube, and place the column part with the lid open on a clean ‘Kimwipe’. Place on a 60 C° heatblock for ~10 min to evaporate residual ethanol. Avoid touching the ‘nipple’ at the bottom of the column with your fingers.
13. Place column into an fresh Eppi tube, and add 30 μl of buffer EB to each column. (ensure EB goes onto the membrane at the bottom, not the wall). Allow to sit for 2 min. Centrifuge 1 min at ~10,000 g.
14. Digest 2 μl of the plasmid preparation with an appropriate restriction enzyme (eg. an enzyme that cuts once or twice) (see restriction digest protocol), and run on an agarose gel to evaluate yield. Note that the Nanodrop may not be accurate for plasmid preps.

#### Adaption for 50ml culture

1. Inoculate 50 ml of LB broth containing the appropriate antibiotic with a large loopful of your E.coli culture from a fresh plate of the same medium.Grow overnight (16-24 h) at 37°C with shaking.
2. Aseptically transfer (pour) the cells into a 50 ml Falcon tube. Find or make a balance tube with the same volume of water. Estimating equal volumes by eye (+/- 1 ml) is OK for low speed centrifuges, but a balance should be used for more sensitive machines or when spinning at higher speeds (>3000g) (for these, need accuracy +/- 0.1 mg).
3. Centrifuge at 4000 rpm for 10 minutes. Pour of supernatant and keep pellet
4. Resuspend the cell pellet in 800 μl buffer P1 by vortexing. Transfer 200μl of the resuspended cells into 4 fresh Eppi tubes. Continue as for 5mL protocol from step 5.

#### Plasmid Prep Solutions

* Buffer P1: 5 mM EDTA, pH 8. Sterilise by autoclaving. Then add RNAse to 300 μg/ml.

Usually this would be prepared by starting with a 0.5 M EDTA, pH 8 stock, and a 10 mg/ml aliquot of boiled RNAse.

* Buffer P2: 1% SDS, 0.2 M NaOH. Not sterilised.

Prepare from stocks of 10% and 2 M NaOH (these don’t need to be sterilised, but it is not a bad idea to prepare these and the final buffer in sterile bottles or tubes containing the correct amount of autoclaved RO water). Keep in a tube or bottle with minimal headspace and tightly closed. Will eventually develop precipitate of Na2CO3 – remake the solution if you see this. Recommend to make fresh approx. monthly.

* Buffer N3: 4.2 M guanidine HCl, 0.9 M K-acetate, pH 4.8. Not sterilised. Adjust the pH using glacial acetic acid
* Buffer PB: 5 M guanidine HCl, 30% isopropanol. Not sterilised. Prepare in a sterile bottle or tube containing the correct amount of autoclaved RO water.
* Buffer PE: 10 mM Tris-HCl, pH 8, 80% ethanol. Not sterilised. Prepare in a sterile bottle or tube containing the correct amount of autoclaved RO water.
* Buffer EB: 10 mM Tris-HCl, pH 8. Autoclaved. Usually this would be prepared by diluting a 1M Tris-HCL pH 8 stock.

Polymerase chain reaction (PCR)

Notes: As with all molecular biology and microbiology work, aseptic technique is crucial. Clean your bench, tube racks, and pipettes with 80% ethanol before starting, and work carefully to ensure you don’t contaminate the PCR reagents or mixtures with microbes, DNA, or enzymes from your hands or the bench or the ice bucket etc etc. Ensure pipette tips, Eppi tubes and PCR tubes are sterile.

This protocol can be used to add restriction sites to ends of DNA sequences. When making primer, add the restriction site sequence to the 5’ end of the primer.

Different PCRs behave differently. Some PCRs are very straightforward while others are very challenging. The factors impacting the difficulty of PCR include the following:

* Type of DNA Template: the easiest template is a plasmid or PCR product containing your sequence of interest. These very clean and have a very high ratio of target sequence to non-target sequence. A medium difficulty template would be genomic DNA from a single microbe.
* Purity of DNA template: highly purified DNA will amplify much better than crude DNA (e.g. boiled cells), which will amplify better than whole cells, which will amplify better than a complex and dirty template (e.g whole soil) (this is unlikely to work at all!). That said, you can certainly do PCR on whole cells of most gram-negative bacteria, including E. coli (this is known as ‘colony PCR’), and this is very useful for rapidly screening clones.
* Type of primers: specific primers work well, degenerate primers work less well, and the higher the degeneracy, the worse they perform. Primers that are designed well (no dimers/hairpins or only weak dimers/hairpins) will work better than primers that are poorly designed (strong dimers/hairpins). Primers that have lots of junk at the 5’ end like long non-target sequences or fluorochromes will perform worse than primers that are exactly the same as the template. Primers with mismatches to the template may still work, but the more mismatches present, the worse they will work, and the closer the mismatches are to the 3’ end, the more serious the problems will be.
* Type of target gene, and its copy number: If your target gene is on a plasmid, it will amplify more easily than a chromosomal gene. If your gene has many copies in the template DNA, it will amplify better than a gene which is very rare. If your target gene is 16S rDNA or some other highly-conserved gene, and your primers are ‘universal’ primers, you might expect problems with contamination, since this gene is \*everywhere\*, including on your hands, on the bench, in your pipette etc. (the negative control with no DNA added is critical in this case!). On the other hand, if your target gene is only found in your particular favourite and unusual organism, then contamination will be less of a concern.
* Size of PCR product: The smaller the product, the easier the PCR. Product size should not cause problems up to ~ 1 kb, but as you go larger than this, the PCR will become increasingly challenging. This also depends a lot on the type of polymerase used (Phusion is better than Pfu, which is better than Taq, in terms of getting large products). Anything larger than ~5 kb is going to be difficult to amplify, even for an experienced user – success here requires using a fancy polymerase, \*excellent\* primers (well-designed, no dimers/hairpins), clean and high-quality template with high signal:noise ratio, fresh reagents, and excellent hands-on technique. In theory, PCR products up to ~20 kb are possible, but these are extremely difficult to obtain in practice.

#### Protocol for setup of PCRs

Notes: The usual PCR is 25 μl volume, this is appropriate for “screening” purposes. However, if you are trying to make a lot of PCR product for cloning, then scale reactions up to 50 μl per tube. A PCR that is working well should give you about 50 ng of product per μl of reaction, thus 2.5 μg per 50 μl. For a cloning experiment, if you prepare 8 x 50 μl PCRs, this should give you plenty of insert DNA (10-20 μg), even if your PCR is not working at high efficiency. This is way in excess of the theoretical requirement, but allows for losses due to subsequent purification steps, and the likelihood that you may need to repeat the experiment a few times

##### Example Reaction Mix (per 25μL)

* 10xbuffer: 2.5μl (->1x)
* sterile milliQ water: 20 μl
* dNTPs (10 mM): 0.5 μl
* primer #1 (50 μM): 0.25 μl
* primer #2 (50 μM): 0.25 μl
* polymerase\* (5 U/μl): 0.25 μl
* DNA template: 1uL

##### For 8 Reactions\*\* (one strip of tubes)

* 10xbuffer: 22.5μl
* sterile milliQ water: 182 μl
* dNTPs (10 mM): 4.5 μl
* primer #1 (50 μM): 2.3 μl
* primer #2 (50 μM): 2.3 μl
* polymerase\* (5 U/μl): 2.3μl
* DNA template : 8 x 1uL

\* This means thermostable DNA pol e.g. Taq, Pfu, Phusion, Q5, etc. but NOT Klenow or T4 pol.

\*\* Calculations done as if 9 x 25 μl reactions were being prepared, to ensure mix doesn’t run out.

Template considerations: The correct amount of DNA template depends on the type of sample. If you are using a plasmid, you only need a tiny bit (~1 ng) because the ratio of target:non-target sequences is very high. For genomic DNA of a single microbe, a more usual amount would be 10-20 ng per reaction. For metagenomic DNA, you may need to go even higher (100 ng per reaction), although note that there are diminishing returns here due to the presence of PCR inhibitors in many templates that will interfere with PCR. For complex templates like soil DNA, the optimum concentration of template often needs to be determined experimentally. More is not always better; in some cases, diluting the template (e.g. 1/10) will yield a band when undiluted template gives no band at all.

##### Set up of Mastermix and Reactions

1. Exact setup of PCR depends on which element is the variable in the reaction. Usually this is the DNA template, but sometimes it is the primer. The ‘Master Mix’ should be made to include everything except the variable being tested. The below protocol and above recipes are written assuming that this variable is the DNA; you need to modify if the same template is being tested with multiple primers.
2. Calculate how many PCRs you need altogether, and thus the total volume of master mix required. It’s important to make more than you need (~10% more), as pipetting errors will always change the expected volumes a little bit. Write up the exact recipe that you need in your lab book.
3. Label your strip tubes on the side, near the top. If you label the lids or the bottom of the tube, the labels tend to come off during thermocycling. Place the tubes in a tube-rack (a 96-well micro-titre plate works well). Leave the lids off the tubes – you are more likely to contaminate the reactions via excessive opening and closing of tubes than from stuff falling in from the air.
4. Retrieve the PCR reagents from the -20°C freezer and thaw them (except polymerase, this remains liquid even at -20°C due to glycerol in buffer). It’s important that the 10x buffer, dNTPs, and primers are thoroughly thawed out before use. Give them a brief vortex or flick to mix. These reagents don’t need to be kept on ice during the time needed to set up the PCR, but don’t leave them on the bench longer than needed.
5. Prepare the master mix in a sterile Eppi tube, containing everything except the variable being tested. Add ingredients in the order listed in the recipes above. Some folks will insist that this must be done on ice, opinions differ. It is true that you will minimise premature polymerase activity by setting up on ice, but whether this really makes a difference for most PCRs is arguable. Setup on ice is supposedly more important for Pfu and Phusion which contain 3’-5’ exonuclease activity which can ‘eat’ your primers during setup. If you set up on ice, risks of contamination from the ice itself must be managed.
6. Aliquot 25uL of the master mix into each of the PCR tubes.
7. Adding the DNA

1. (DNA template in solution) Add DNA template (1 μl) to all the tubes, being very careful to match the sample #’s to the labels on the tubes. You need to concentrate your attention here and really focus in order to avoid missing a tube or putting two templates in the same tube. Be careful to change tips between every DNA template.
2. (colony PCR: DNA in cells). Touch a white tip onto a colony or patch of growth and pick up a \*small\* amount of cells. It should be just enough so that you can see there are cells there, not much more than this. Then dip the tip in and out of the master mix in the first PCR tube approx 5 times and use the same tip to streak out a patch on a patch plate. You don’t need to wipe the whole chunk of growth on the inside of the tube, just the dipping in and out action will dislodge enough cells to give you enough template DNA. Less is more in this case! Repeat for other colonies/patches.

8. Put lids on tubes, ensure they are snapped on tight, place immediately in thermocycler. Double check your program parameters before starting. See below for detailed thermocycling instructions.
9. Return all reagents to the freezer.
10. After thermocycling is completed, run out your PCRs on an agarose gel to check their size and yield. Typically, we would load 5 μl of PCR mixture. This should give a very strong band if the PCR has been successful.
11. If you are intending to use the PCR product for cloning, check for the presence of non-specific bands (usually fainter, smaller bands than the expected one) – if these are numerous and/or strong, you may need to try a higher annealing temp, or revisit primer design, before the product can be cloned. Alternatively, you can cut out the desired band from the agarose (see our Purification of DNA via spin column protocol).

#### Thermocycling

A standard thermocycling protocol is given below (for Taq polymerase)

* Initial denaturation: 95°C, 5 min
* Denaturation: 95°C, 30 sec
* Annealing: 5°C lower than the melting temperature of the primers, 30 sec
* Extension: 72°C, 1 min per 1000kb product (differs between polymerases)
* 35 cycles of these steps
* Final extension: 72°C, 10 min
* Hold: 15°C

The numbers that are in bold indicate variables that need to be optimised for every individual PCR. Sometimes, other variables may also need to be modified too, as below
Annealing temperature: This is typically set at 5°C lower than the melting temperature (Tm) of the primers. If these Tm’s are different, use the lower one for this calculation. This calculation is very crude, and it is best to optimise annealing temperature experimentally by doing a ‘gradient PCR’, e.g. testing an annealing temp range from 5° lower to 5° higher than the predicted best temp. Aim to make primers with Tm’s around 65°C, which gives a predicted annealing temp around 60°C. The usable range for annealing temp’s is approx 50°C – 70°C.

Extension time: This is set primarily by the length of the desired PCR product. For Taq polymerase, this is 1 minute of extension per kb of product. However, note that different polymerases require different extension times: Pfu needs 1.5 min per kb, and Phusion needs 0.5 min per kb. If you use a longer extension time than necessary, you risk increasing the yield of non-specific products; if you use a shorter extension time than necessary, you risk a low yield of the desired product.
Denaturation: The optimal denaturation conditions for Phusion pol are different than for Taq or Pfu. Change the initial denaturation to 98°C - 30 sec, and the denaturation in each cycle to 98°C - 10 sec.

Number of cycles: This can be increased to 35 or even 40 cycles to give a higher yield of product, but this risks introducing more mutations into the PCR amplicons, and also gives a higher chance of secondary non-specific products being formed. Conversely, the # of cycles can be reduced to 25, which will minimise mutations and non-specific products, but will also lower yields.

Hold temperature: It’s OK to leave your PCRs in the machine overnight. While some protocols recommend fridge temp (4°C) for this ‘Hold’ step, this is actually bad for the machine – it will accumulate condensation on the block, which will degrade the block over time. A good compromise is to set the hold temp to 15°C, which keeps the samples cool(-ish), maintaining them in good condition, but is not cool enough to cause condensation to form.

PCR Enhancer Additions: Two enhancers that might be worth testing for problematic PCRs are Bovine Serum Albumin (BSA) and dimethylsulfoxide (DMSO). BSA is good for binding inhibitors and is typically added at 0.5 mg/ml final conc. from a 10 mg/ml stock (=1.25 μl per 25 μl reaction). DMSO is good for enabling amplification of GC-rich templates and is typically added at 1 μl of neat DMSO per 25 μl reaction (=4%).

Colony Screening PCR

![](https://static.igem.org/mediawiki/2019/1/12/T--Sydney_Australia--ProtocolsScreeningPCRPrimers.png)

#### Prepping your Primers

This diagram shows some of the basic
PCRs that can be used to determine
whether a plasmid clone is:

* The desired recombinant
* Some other kind of recombinant
* Vector only

![]()

PCR using primers F1-R1 is often the
starting point for screening. This is
known as  left junction PCR.  Getting a
product of the expected size in this PCR
is a good indication you have the right
recombinants.

PCR using primers F2-R2 (right junction PCR) can then be done on colonies positive in
left junction PCR. If the clones are also positive in this PCR, it is further evidence that
they are the construct you’re after.

PCR using the primers F1-R2 is known as a  spanning PCR or an insert spanning PCR.
This PCR will amplify the entire insert region. This PCR can be used to determine if the
insert is the correct size, but bear in mind that this is  not an all or nothing PCR like the
junction PCRs. In spanning PCR the clones will give a small product if they have no
insert, and a large product if there is an insert. However, if you have a mixed
population of cells you may get a false negative result as the small product will be
amplified preferentially, even if there are positive clones in there too. This is why pure
colony isolates are so important. Spanning PCR is most often used to prepare the
insert region for sequencing rather than initial clone screening as longer PCR
products take longer to thermocycle.

Primers used for the amplification of the original DNA insert (not shown in the
diagram) can also be used for screening, but they will not tell you whether the insert
is oriented correctly like the junction PCRs will.

Positioning of the primers is important, especially for later steps such as sequencing.
Make sure that all primers (F1, F2, R1, R2) are at least 50bp away from the ligation join
- as the first 20-30bp of sequence read are usually ‘junk’. However, if the primers are
too far away from the ligation join you’re amplifying and sequencing things that you
don’t need to, which increases time and expense. Placing the primers about 100-150bp
away from the ligation join is good, and ought to give a positive PCR product of
around 200-300bp.

#### Initial Patching and PCR Screening

1. Choose 20 well-isolated colonies from your transformation plates. (if your
   construct has colour selection, ensure you pick colonies that are not connected to
   any negatively coloured colonies). Circle and number these on the base of the
   plate.
![](https://static.igem.org/mediawiki/2019/1/1f/T--Sydney_Australia--ProtocolsScreeningPCRPatchPlate.png)2. Label three PCR strip tubes with numbers 1-20, leaving
   three blanks (one per strip) as negative controls.
3. Using an agar plate containing the appropriate
   antibiotic, divide the plate into patches. On the
   bottom of the plate draw a 5x5 grid. Label 20 of the
   squares with numbers 1 to 20 (see diagram).
4. Prepare a PCR master mix with appropriate left
   junction primers, enough for ~30 PCRs. Aliquot this
   into 3 x 8 lots of 25uL in the labelled PCR strip tubes.
   Taq polymerase is good to use for this job as it is very
   robust to any ‘mess’ in the PCR, but other polymerases
   can also be used.
5. Using a white pipette tip (suitable for p10 or p2 pipettes),
   pick up some growth from one of your colonies of interest. Make
   sure you can see some cells on the tip, but you don’t need a lot.
6. Dip the tip in and out of the first PCR tube five times. You don’t need to dislodge
   the whole chunk of growth, enough cells will fall into the mix to give you a product.
7. With the same tip, transfer the remaining growth onto the first patch on your
   agar plate by scratching the tip across the surface of the agar to make a set of
   three close parallel lines (see diagram). Stay clear of the borders of each square,
   we don't want the colonies to get too friendly with each other!
8. Repeat steps 5 - 7 for the remaining colonies, making sure to be careful that you
   match your tube number to the plate square number each time. Cross of circled
   colonies after you have sampled them - no need to test one colony twice!
9. Put the PCR tubes in the thermocycler and incubate the agar plate overnight.
10. Run the PCR products on an agarose gel and note which colony numbers gave a
    positive result.
11. If none of your clones are positive then you may need to test another twenty from
    your transformation plate. However, if none of those are positive either you will
    need to do some troubleshooting as something is likely wrong - check your
    ligation and transformation controls, check your primer sequences for issues such
    as dimers, and make sure your thermocycler conditions are appropriate for your
    polymerase and your primers. You may end up needing to repeat the cloning
    process. If a second cloning still yields no positive colonies it is possible that your
    gene of interest is toxic to your host cells - you may need a diferent vector (e.g.
    inducible, lower copy number) or a diferent host or diferent incubation
    conditions.

#### Secondary PCR Screening

1. After getting between three and seven positive colonies from your primary
   screening you may want to confirm these results with a right junction PCR.
2. Set up a PCR master mix with appropriate right junction primers. Make enough
   master mix so there is at least 10% more than you need so you don’t run out.
3. Make up another agar plate, but this time with just enough squares for the
   number of positive colonies. It may be helpful to you to label the squares with the
   numbers of the positive patches (i.e. if your positives were numbers 7, 12, and 19
   you would label your three squares with those numbers rather than 1, 2, and 3.)
4. Follow steps 5 - 8 from the above protocol, taking some colonies from the patch
   plate you made.
5. Put the PCR tubes in the thermocycler and incubate the agar plate overnight.
6. After thermocycling, run the products on an agarose gel. Note which colonies
   were positive.
7. Choose one positive colony to inoculate a 100mL LB-antibiotic broth to prepare for
   a plasmid prep.
8. Keep the patch plate! Wrap it with parafilm and keep it in the cool room. You may
   end up needing to use a diferent positive colony if there are problems with the
   first one.

Even though you’ve performed two junction PCRs you can’t be sure your construct is
completely correct. It is advised that you validate your construct using a restriction
digest  or  through sequencing . Small mutations cannot be easily detected by junction
PCR, and they can completely change the functionality of your construct!

Diagnostic restriction digest for plasmids

1. Choose one or a combination of restriction enzymes that will cut the plasmid 1-3 times. Make sure that all the predicted fragment sizes are different enough to separate on a gel. If you’re using multiple restriction enzymes, make sure that they all work in the same buffer and at the same temperature.
2. Get your buffer out of the freezer and ensure it is totally thawed and mixed before use.
3. Get restriction enzymes out of freezer and put them on ice immediately. It is critical that the enzymes stay cold at all times.
4. Set up the digest in a sterile Eppendorf tube in the following order:

2uL 10x buffer

YuL MilliQ water

XuL plasmid, containing 250ng DNA

1uL each RE

----------------------------------------------

20uL total

Ensure you are changing tips between every addition to avoid contaminating the restriction enzymes.

Ideally, you would not use more than 5uL of plasmid. If you use more that 5uL, the digest may not work due to junk in the plasmid prep (e.g. salts) interfering – in this case, consider further purification and concentration of the plasmid e.g. by ethanol precipitation. Calculate the amount of MQW needed (Y) by subtracting the total volume, 20uL, from the amount of plasmid needed (X) and the number of restriction enzymes being used. - Incubate at the preferred temperature. If using HF enzymes, 15 minutes is sufficient, but can digest for up to 24hrs. 30 minutes – 1 hour is optimal.
- Run on agarose gel to check. Note: For Bam restriction enzymes, you must use SDS containing loading buffer or purify or heat kill the sample (15 min 80°C) before running the gel, as Bam binds tightly to the DNA and will slow the migration of the DNA and change the apparent size of the bands.

Mung Bean Nuclease End Blunting

This protocol is to turn the sticky ends left over from a restriction digest into blunt ends. Mung bean nuclease has the ability to chew back the overhang.

Can be done in the same reaction mixture as the digest that creates sticky ends or can be done subsequently after heat-killing or spin column purifying DNA. Reaction can be scaled up or down as needed.

A reaction contains:

* 10x Cutsmart Buffer
* A certain number of ng DNA (make sure you know how many!)
* 1 Unit Mung Bean Nuclease per ng DNA. The concentration of NEB Mung Bean Nuclease is 10,000U/mL, which is the equivalent of 10U/uL. Therefore, the number of uL of nuclease is the number of ng of DNA divided by 10.
* Total to desired volume with MQW (30uL is a good size, depends on the amount of DNA)

Make up reaction in Eppendorf tube. Incubate at 30°C for 30 minutes.

After the 30 minutes is over, purify the DNA by spin column. DO NOT ATTEMPT TO HEAT KILL!

Making and Running Agarose Gels

Making gels is one of the most crucial parts of any iGEM project. They can be used for gel purification, restriction digest diagnostics, and so much more. Here's our preferred method for making gels:

1. Figure out how many samples you have and choose well-formers and gel volume accordingly: the smallest gel tank uses 50 ml agarose and takes up to 10 samples, the largest tank uses 150 ml agarose and takes up to ~40 samples (if two rows of well-formers used). Remember to leave a lane for the molecular weight marker!
2. Decide what agarose concentration and what buffer are appropriate. For many gels, 1% agarose and 1x TAE will be fine. If you are trying to separate very small DNAs (<500 bp), you can increase up to 2% agarose, and if you are trying to separate large DNAs (>5 kb), you can decrease down to 0.7% agarose. If you want to run a gel very quickly (15mins), use 1 x LAB buffer. If you need to purify DNA from the gel for cloning or sequencing, use TAE (borate from TBE or LAB interferes with later enzyme reactions).
3. Decide whether you need very accurate band-sizing or not. If you do not (e.g. just want to see band or no band when screening clones), you can put the detection dye (GelGreen or Thizole Orange) into the gel. If you do need high accuracy, instead use post-staining.
4. Set up the gel-casting tray by masking-taping the ends of the tray. Place the well-former (comb) such that it is level and straight and sitting ~2 mm above the base of the gel forming unit. It is crucial that the well formers are NOT TOUCHING the casting tray – in that case, your wells will have holes in the bottom and you will lose your samples.
   5. Using a small, clean, dry conical flask, weigh out the appropriate amount of agarose powder for the correct and % agarose gel that you intend to make, then pour in the appropriate volume of TBE or TAE buffer. Put in microwave, and heat on high for 1 minute. Put on heat resistant gloves, then take out to examine whether all the agarose is dissolved. If not, replace for another 30 sec on high power, and repeat examination and heating until all dissolved.
5. Cool the conical flask down under a stream of cold water, while swirling. Be careful not to get water INTO the flask. After about 20 seconds, see if you can hold the flask comfortably in your hand (no glove) – if it is too hot to hold, continue cooling, re-check every 20 seconds or so. When the flask feels warm but not uncomfortably hot, it is ready to pour (this is approx. 45-50°C). If you cool it too much, it may set solid. In this case, microwave again (as above) in 30 sec increments to re-dissolve.
6. If appropriate: add GelGreen or Thizole Orange dye to the gel. For gels with long run times, >40mins, do not use Thizole Orange, as it will run off the gel. Use 5uL per 50mL gel of stock for both dyes.
7. Pour the molten agarose into the casting tray. Use a single smooth motion, don’t stop and start. Stop when the agarose is 3/4 of the height of the ‘teeth’ on the well-forming comb. Or alternatively, stop when you judge the wells are deep enough to hold the amount of sample that you need to load. What you DON’T want to do is over-fill the agarose so that it goes over the teeth – this will result in a channel that connects all the wells, and subsequent cross-contamination of the samples.
8. Allow the gel to set. This takes about 15 minutes but will be faster if set up in the cold room.
9. When gel is set, remove the masking tape, and put the gel in the electrophoresis tank. Fill the tank with enough of the appropriate buffer to fills both wells and covers the gel by 1-2mm, then carefully pull out the well formers (straight up, don’t yank them side to side or forwards and back). The reason for the buffer is to stop the wells collapsing on themselves (this can happen with thin wells at lower % agarose).
10. Choose the appropriate molecular weight marker. For DNAs <1 kb, use the NEB 100 bp ladder. For DNAs from 1 kb – 10 kb use the NEB 1 kb ladder. You may need to load two different ladders, one on each side of the gel.
11. Prepare a row of spots of purple loading dye on a Parafilm strip, of appropriate volume and number. This is a 6x buffer.
12. List in your lab book a map of which samples will be loaded in which locations. Avoid the first and last wells if possible, these tend not to run as straight as the more central wells.
13. Mix your first sample with the first spot of loading dye, and load into the first well. Steady the pipette tip with the finger of your non-pipetting hand to ensure accurate dispensing. The tip needs to be just inside the well, don’t push it all the way down in the well. Change tips, mix up the next sample with blue dye, and load again. Repeat for all samples, including the marker.
14. Put the lid on the gel box, check that the terminals are connected correctly (negative terminal should be closest to the wells, positive terminal is far from the wells, ie. Run towards Red). Run the gel at ~150 volts (small gel box) up to ~250 V (large gel box). Check that you can see gas bubbles at the electrodes when you start running the gel. If not, check all the wire connections.
15. Stop the gel when the fast-running blue dye (bromophenol blue) (= usually the only blue dye) is near the end of the gel (this may take 30-60 min). You may need to run longer to get good separation of large products (>5 kb). Run for shorter time for small products (<500 bp), or they may run off gel.
16. If you added GelGreen or Thizole orange to gel, it is now ready to image on the transilluminator. Otherwise, post-stain in a solution of Thizole Orange (5 μl per 50ml water) for 30 min on a mechanical rocker, before transilluminating.

Purification of DNA via spin column (for DNA in solution)

#### GENERAL NOTES

This protocol is good for restriction fragments, small digested plasmids, and PCR products. It is NOT
good for genomic or chromosomal DNA. Use the FastPrep reagents or CTAB-phenol type prep
instead for genomic DNA. Digested plasmids work well up to about 5 kb, but the yield quickly falls off
past this point. Generally, plasmids don’t purify as well as PCR products on these columns.

1. Figure out what volume of sample you have. If this is less than 100 µl, then make it up to 100 µl
   with TE. The volume you now have will be called “1 volume”. Add 5 volumes of buffer PB to the
   DNA solution. e.g. if you have a 100 µl restriction digest, add 500 µl PB.
2. Place a silica-based spin column (e.g. ‘Econospin’) into its 2 ml catch tube (if it isn’t already set up
   that way). Load up to 750 µl of the DNA-PB mixture onto the column. Spin at ~10,000 g for 30 sec.
   Discard the flow-through into culture waste.
3. If you still have more DNA-PB mixture left, repeat the previous step until all of the mixture has
   been put thru the column. The columns will hold a total of ~10 µg DNA, which is a lot!
4. Add 750 µl of buffer PE to the column, spin ~10,000 g for 30 sec, discard flow-through.
5. Repeat step 4.
6. Spin again for 30 sec to remove all traces of PE from the column. Discard both the flow-through
   and catch tube, and transfer the spin column onto a clean Kimwipe. Leave the column lid open.
   Transfer Kimwipe to 60°C oven, and allow to dry for 10 min.
7. Transfer spin column to a sterile 1.5 ml Eppi tube, and add 20-50 µl\* of EB buffer to the centre of
   the spin column – ie on the membrane, not the walls of tube. Allow to sit for 2 min. Spin at ~10,000
   g for 1 min, retain Eppi tube with DNA solution in EB, discard spin column.

\* Usually we want high concentration rather than high yield, so use 20 µl. If max. yield is important,
or if you have lots of DNA, use 50 µl. Note that you lose approx 5 µl EB during the procedure.

- Buffer PB: 5 M guanidine HCl, 30% isopropanol. Not sterilised. Prepare in a sterile bottle or tube containing the correct amount of autoclaved RO water.
- Buffer PE: 10 mM Tris-HCl, pH 8, 80% ethanol. Not sterilised. Prepare in a sterile bottle or tube containing the correct amount of autoclaved RO water.
- Buffer EB: 10 mM Tris-HCl, pH 8. Autoclaved. Usually this would be prepared by diluting a 1M Tris-HCL pH 8 stock.

Purification of DNA via spin column (for DNA in a gel slice)

#### General Notes

This protocol is good for restriction fragments, small digested plasmids, and PCR products. It is NOT good for genomic or chromosomal DNA. Digested plasmids work well up to about 5 kb, but the yield quickly falls off past this point. Generally, plasmids don’t purify as well as PCR products on these columns.

Gel-purification: Don’t expose the DNA to any UV light and don’t use borate-containing buffers in the gel. Even minimal exposure to short-wave UV light will greatly damage DNA, and make it hard to ligate later. Long-wave UV (e.g. hand-held lamp) is less damaging than short-wave UV (transilluminator), but best to avoid UV altogether. Borate can cause problems for later enzyme steps.

#### Notes on Gel Extraction Using Gel-green (recommended)

1. f you need to purify DNA from an agarose gel, use TAE-agarose (don’t use TBE!). Put Gel-Green (1 μl) in the gel. Load plenty of DNA so it is easy to visualize.
2. Run gel as usual, then visualize under blue illumination (NOT UV!) in a dark room or box. There is a blue light in the lab for this purpose and orange-tinted glasses also help. Cut out your band(s) of interest with a clean, sharp scalpel blade, and put them into an Eppi tube.
3. Weigh the tube containing gel slice and subtract weight of empty Eppi tube to figure out the weight of gel. Proceed to ‘main protocol’ below.
4. Weigh the tube containing gel slice and subtract weight of empty Eppi tube to figure out the weight of gel. Proceed to ‘main protocol’ below.

#### Main Protocol

Be sure to have your gloves and safety glasses on. QG contains guanidine which is somewhat toxic.

1. If your weight of agarose is less than 100 mg (=100 μl), then make it up to 100 μl with TE. The volume you now have will be called “1 volume”. Then add “3.5 volumes” of QG buffer to your gel slice. eg. if you have 100 mg of agarose, add 350 μl of QG.
2. Slice up or mash the agarose in an Eppi (<300 mg agarose) or a McCartney bottle (>300 mg). This will speed dissolution. Melt the agarose with heating (60°C for 5 min is usually enough, mix occasionally), then when it’s all dissolved, allow to cool to room temp.
3. Add “1.5 volumes” of isopropanol to the mixture. e.g. for 100 mg agarose, use 150 μl isopropanol.
4. Place a silica-based spin column into its 2 ml catch tube (if it isn’t already set up that way). Load up to 750 μl of the DNA / QG / isopropanol mixture onto the column. Spin at ~10,000 g for 30 sec. Discard the flow-through into culture waste.
5. If you still have more DNA /QG / isopropanol mixture left, repeat the previous step until all of the mixture has been put through the column. The columns will hold a total of ~10 μg DNA, which is a lot!
6. Add 750 μl of buffer PE to the column, spin ~10,000 g for 30 sec, discard flow-through.
7. Repeat step 6.
8. Spin again for 30 sec to remove all traces of PE from the column. Discard both the flow-through and catch tube, and transfer the spin column onto a clean Kimwipe. Leave the column lid open. Transfer Kimwipe to 60°C oven, and allow to dry for 10 min.
9. Transfer spin column to a sterile 1.5 ml Eppi tube, and add 20-50 μl\* of EB buffer (5 mM Tris, pH 8) to the centre of the spin column – ie on the membrane, not the walls of tube. Allow to sit for 2 min. Spin at ~10,000 g for 1 min, retain Eppi tube with DNA solution in EB, discard spin column.

\* Usually we want high concentration rather than high yield, so use 20 μl. If max. yield is important, or if you have lots of DNA, use 50 μl. Note that you lose approx 5 μl EB during the procedure.

#### DNA Purification Solutions (Spin Colummn)

* Buffer QG: 5M guanidine HCl. Not sterilized. Prepare by adding GuHCl to autoclaved RO water in a sterile bottle/tube.
* Buffer PE: 10 mM Tris-HCl, pH 8, 80% ethanol. Not sterilised.

Add appropriate vol. of autoclaved 1 M Tris-HCl to the correct amount of autoclaved RO water in a sterile bottle or tube, then add 100% ethanol to give 80% final conc.

* Buffer EB: 10 mM Tris-HCl, pH 8. Autoclaved. Add appropriate vol. of autoclaved 1 M Tris-HCl to the correct amount of autoclaved RO water in a sterile bottle or tube.

Preparation of Chemically
Competent Cells Using
Rubidium Chloride

Use excellent aseptic technique at all times. All materials must be sterile.
Protocol can be scaled up or scaled down as required. 100mL E. coli culture produces
about 50 x 220uL aliquots of competent cells. Each aliquot is sufficient for 4
transformations, with a little bufer for safety.
Competent cells can be used fresh, so feel free to make them fresh for your
experiment but as this is quite an involved process, do your labmates a solid and
make a big batch to put into the freezer for future use.
Freezer stocks will not survive multiple freeze-thaw cycles so once you take an
aliquot out of the freezer any leftovers must be disposed of.

After step 7 you will need to work on ice and in the cold room to increase the quality
of the final cell prep. These steps will be written in blue for easy differentiation.

####

1. Streak desired E. coli strain (e.g. TOP10, BL21(DE3), JM109) from glycerol stock or
   other source onto plain LB agar, no antibiotics (means you need to be extra
   careful!). Incubate overnight at 37C.
2. Check that your culture from the night before looks pure. (hint: if it’s green and
   fuzzy, that aint E. coli). If it’s looking good, inoculate a plain 5mL LB broth with
   several of the colonies from your LB plate. Incubate overnight (16-24hr) at 37C
   with shaking.
3. At the beginning of the day, place two centrifuge tubes in the freezer, ready to
   use.
4. Aseptically inoculate 100mL plain LB broth (in a 500mL Schott bottle or
   Erlenmeyer flask) with 3 ml of overnight culture; this should give an initial OD600
   of ~0.05. (check this to be sure).
5. Grow cells at 37C with shaking until the culture reaches an OD600 of ~0.5 (if you’re
   short on time, above 0.3 is passable. If you overshot it, anything up to 0.7 is okay).
   This can take up to 3 hours, but some E. coli are over achievers so after the first
   hour check the OD every 15-30mins.
6. Aseptically transfer (also known as carefully pour) the cells into the cold, sterile
   centrifuge tubes or bottles (one 250 ml Sorvall bottle or 2 x 50 ml Falcon tubes)
   you prepared earlier. Balance these, either by transferring culture aseptically
   between the tubes/bottles or by adding sterile water or LB to the lighter tube/
   bottle.
- (Brrr… now’s the chilly part!) Centrifuge at 4000 rpm (~3000 g in Centaur/
  Centurion machine) at 4°C for 10 minutes. Note: centrifuge needs to be cold. Turn
  on and set temperature beforehand. You can spin faster, up to say 7000 g, but
  above this point, faster is not better, and cell pellets will be hard to resuspend, but
  if you skipped arm day by all means give it your best shot.
- Working in the cold room with the cells on ice, pour of the supernatant into
  culture waste (don’t let the centrifuge tube/bottle actually touch the edge of the
  culture waste bottle). Try to remove as much of the liquid as possible – give it a
  shake / tap to assist this.
- Resuspend the pellet gently in 33mL RF1 solution by vortexing and/or shaking the
  tube/bottle. Its OK to be rough with the cells at this stage in the process, but you
  shouldn’t need to shake or vortex for more than 10 sec or so.
- Incubate on ice for 1 hour, then pellet the bacteria again at 4000rpm, 4°C for 10
  minutes.
- Working in the cold room, pour of the supernatant into culture waste. As before,
  try to remove as much of the residual liquid as possible.
- Resuspend the pellet in 8mL RF2 solution by vortexing or shaking. At this stage,
  the cells have become more fragile due to the RF1 treatment, so its important not
  to shake/vortex any longer than ~10 sec (this shouldn’t be necessary).
- . Incubate on ice for 15 minutes. While the cells are incubating, set up all your Eppi
  tubes (~50) on ice with the lids open, so they are pre-chilled, and ready to receive
  cells. Label these tubes on top with the strain name before putting them on ice
  (labelling becomes difcult with cold and wet tubes!). Be careful to only push the
  tubes only about 2/3 of the way into the ice. If they are pushed in too far (right up
  to the lip of the tube), you risk getting ice or melted ice (not sterile!) into your cell
  aliquots
- Working quickly (but still carefully!), aliquot 220 µL of cell suspension into the prechilled Eppi tubes. Once dispensed, close tube lids tightly, and collect all tubes into
  a bag/box with a clear and prominent label and store it immediately in the -80°C
  freezer. It’s a good idea to label both the outside of the bag/box AND place a
  label written on paper inside. Label with the date you made the batch so they can
  be easily identified if anything goes wrong in testing.
7. Test the transformation efciency of the freshly-prepared competent cells using a
   known amount of a plasmid standard (see protocol for heat shock
   transformation).
8. Streak out a loopful of the cells onto plain LB medium (using either a sample of a
   frozen aliquot or some residual cells remaining in the large centrifuge tube/
   bottle), and incubate at 30°C for three days to allow any of the common types of
   contaminants (e.g. Staphylococcus) to grow. This is to check the purity of the cell
   stock. The streak-plate should look completely uniform, with colonies of only one
   type (E.coli), and no heterogeneity in the initial patch or the streaklines which
   would indicate a mixture of bacteria is present.
9. If either of your tests fail (e.g. you do not get transformed cells, you get
   heterogeneity on your streak plate) dispose of your batch of competent cells.

#### Competent Cell Solutions

##### RF1 Solution

* 500mL Reverse Osmosis Water (ROW)
* 6.06g RbCl (final conc. 100mM)
* 4.95g MnCl2 (final conc. 50mM)
* 1.47g K Acetate (final conc. 30mM)
* 0.74g CaCl2 (final conc. 10mM)
* 75g Glycerol (15% w/v)

##### RF2 Solution

* 500mL Reverse Osmosis Water (ROW)
* 1.06g MOPS (final conc. 10mM)
* 0.6g RbCl (final conc. 10mM)
* CaCl2 (final conc. 75mM)
* 75g Glycerol (15% w/v)

For both solutions: add all the ingredients to 300mL of the 500mL ROW, then make
up the final volume using a measuring cylinder.

For RF1:  After adding all the ingredients, adjust pH to 5.8 with conc. acetic acid. pH will
change very quickly and only requires ~ 10µL of acetic acid. Sterilize by filtration into
an autoclaved media bottle.

For RF2:  After adding all the ingredients, adjust pH to 6.8 with NaOH or HCl, as
appropriate. Sterilize by filtration into an autoclaved media bottle.

Growing E.coli for HIS tag protein purification.

#### Reagents

* Suspension Buffer: 1x TE (10mM Tris, 1mM EDTA pH 8).
* Ni+ bead suspension in 20% EtOH.
* Buffer A: 20 mM Tris, pH 8
* Buffer B: 20 mM Tris, 500 mM NaCl, 5 mM imidazole, 0.1% Triton X-100, pH 8
* Buffer C: 20 mM Tris, 500 mM NaCl, 150 mM imidazole, 0.1% Triton X-100, pH 8
* Buffer D: 50 mM Tris-HCl (pH 7.5), 20% glycerol, 0.1 mM EDTA, 1 mM DTT (Add the DTT after autoclaving.)

#### Growing and collecting cell lysate

1. Inoculate 500mL of LB containing the appropriate antibiotics with your cells contain your HIS tagged constructs. A large loopful of cells from a plate resuspended in 1mL of plain LB or an overnight 5mL seed culture of your cells should be enough of an inoculant.
2. If you are using a T7 polymerase containing cell line (BL21 etc) and vector with a HIS tagged sequence under a T7 promoter incubate your cultures at 37oC with shaking (200rpm) until OD600 0.4-0.6 then induce with IPTG (1mM)
3. Leave the cultures to shake (200rpm) at your desired protein expression temperature overnight. Deciding the ideal expression conditions may require sampling of cell lysates from smaller cultures under various conditions using SDS PAGE. Good working temperatures to test expression include 15oC, Room Temp (21oC -25oC), 30oC and 37oC. See ‘SDS PAGE of Cell Lysates’ protocol for lysate testing methods.
4. Centrifuge the cultures (3900g, 15min, room temp) and pour off the supernatant.
5. Resuspend cell pellet in 25mL of Buffer A by vortexing.
6. Centrifuge once more (as above) and pour off the supernatant.
7. Resuspend the pellet in 5mL of Suspension Buffer.
8. Freeze suspended cells at -30 for 30 minutes.
9. Thaw suspension and add 30uL of 1x protease inhibitor cocktail.
10. Transfer 500uL aliquots of the suspension to sterile bead beating tubes (containing 2 large beads (5mm) two small scoops of medium beads (60 mesh) and two small scoops of small beads (75-150 microns).
11. Bead beat tubes for 30 seconds then move tubes to ice for 2 minutes. if results suggest poor lysis repeat this step.
12. Centrifuge bead beating tubes at 15,000g for 5 minutes (preferably at 4oC)
13. Move tubes back to ice immediately
14. Combine the supernatant from all bead beating tubes into one prechilled 15mL Falcon tube and keep on ice (be careful not to collect any glass beads)
15. Collect 30uL of your now combined cell lysate in a 1.5mL eppendorf labelled ‘Cell Extract’ and freeze at -30oC for future SDS loading (see ‘SDS PAGE of Cell Lysate’ protocol). Proceed with HIS tag purification with the remainder of your cell lysate.

HIS tag protein purification.

1. Pack a 10mL syringe tightly with 1-1.5cm of non absorbent cotton wool (plunge hard to compact the cotton wool at the base of the tube.
2. Suspend your column over ice and remove the plunger and add 2-3mL of your Ni+ bead slurry to the column
3. Allow the beads to settle then slowly add 10mL of Buffer B to the column, allowing the solution to move through and out the bottom of the column into a waste beaker. Ensure the column does not dry out however.
4. Add your cell lysate solution and allow it to move through the column. Collect the flow through in a 15mL falcon tube labelled ‘Flow sample’ on the ice below.
5. Add 20mL of Buffer B to the column gently to flush out any unbound protein in the column. Collect the flow through in a 50mL falcon labelled ‘Wash sample’
6. Add 5mL of Buffer C to the column and collect the flow through.
7. Repeat step 6 twice more collecting the flow through in separate tubes each time.
8. Pipette 30uL of each fraction to new individual 1.5mL eppendorf tubes for SDS PAGE analysis (See ‘SDS PAGE of Cell Lysates’) (These can be frozen at -30oC). If required you store your protein for immediate use at 4oC or freeze them at -30oC. Note that the activity of your enzymes may be affected by long term storage of your enzyme at these conditions.
9. Prepare your ‘Eluted’ fractions for dialysis against 1L of Buffer D overnight at 4oC with a low speed magnetic stirrer.

SDS PAGE of Cell Lysates

#### Preparing Cell Lysates

1. Prepare 1.5-2mL bead beating tubes using the appropriate amount of glass beads. Two 5mm beads (large) and two small scoops of both 60 mesh beads (medium) and 75-150 micron beads (small) should be enough for one tube.
2. Sterilize by autoclaving.
3. Pellet your cells from broth or scrape off a plate and resuspend in 500uL of 1xTE (10mM Tris, 1mM EDTA pH 8) and 5uL of a 1x protease inhibitor cocktail (This will only extract soluble protein from your cells). How much culture to use is tricky to know, more concentrated is usually better, but too many cells and suspension may be gloopy and hard to work with. Use more bead beating tubes if working with larger cultures to avoid this problem. The pellet from 2mL of broth culture or a large loopful from a plate is a good starting point for 1 bead beating tube.

To examine the insoluble fraction of your cells resuspend the culture in 500uL of SDS sample buffer (4% SDS, 20% glycerol, 0.002% Bromophenol Blue, 5% fresh B-mercaptoethanol, 62.5mM Tris-HCL pH6.8).

4. Take your tubes over to the bead beater and unscrew the tube-holder.
5. Space your tubes around the disc (make sure the caps are on tight and the sides of the tubes are labelled!), replace the cover as tight as you can.
6. Beat for 30 seconds. If results suggest poor lysis, or if you are working with Gram-positive bacteria or yeast etc, you will need to do multiple cycles of beating – in these cases, chill your tubes on ice for 1 min between beatings. You may need 5 x 30 sec beatings to fully lyse tough cells like mycobacteria.
7. Centrifuge at ~10,000g for 5 minutes ideally at 4 °C to remove beads and any remaining whole cells. If you must centrifuge at room temperature keep lysates on ice for 5 minutes before centrifuging at ~15,000g for 2 minutes.
8. Retain the supernatant and keep on ice.
9. Check the absorbance of supernatants using a Nanodrop (Protein setting or 280nm wavelength) to determine mg/mL (Abs of 1 = 1mg/mL protein). Ideally we want around 5mg/mL for nice PAGE gels (between 2 and 10mg/mL may be OK). If It’s less than 2mg/mL either repeat with more cells or bead beat for more cycles. If it’s more than 10mg/mL dilute down to 5mg/mL. Keep in mind the Nanodrop reading is only an estimate for protein concentration, since there is other junk in the cell extract that will interfere. For some applications like enzyme assays, you may need to use a different protein quantitation methods.
10. Using your nanodrop estimates calculate how much of each sample you’ll need to get a constant amount of protein in each lane on your PAGE gel. It’s very important to try and match the total protein in each lane so that we can compare e.g induced vs uninduced samples etc. between 25-50ug per lane usually works out well. For example if your sample is 5mg/mL that’s equal to 5mg/uL so for a 40ug lane you would need 8uL of your sample.
11. Add an appropriate amount of SDS-PAGE sample buffer to your sample (final conc: 4% SDS, 20% glycerol, 0.002% Bromophenol Blue, 5% fresh B-mercaptoethanol, 62.5mM Tris-HCL pH6.8). i.e for the 8uL sample above you would need ~8uL of a 2x SDS-PAGE sample buffer stock solution. Make up 50% more sample + SDS sample buffer than you actually need to prevent loss in the next step.
12. Heat samples at 100°C for 3min then allow them to cool down before loading them on a gel.

#### Preparing your SDS PAGE gel.

1. Prepare your 1x SDS running buffer first.

10x SDS running buffer.

##### Reagent

* SDS
* Tris base
* Glycine
* MilliQ

##### Amount

- 10g
- 30.3g
- 144g
- To 1L

pH to 8.5

2. Take your prepared gel cassettes (precast or homecast) and remove the well comb, holders (home cast) and/or bottom tape (precast).
3. Put them into the housing with the short plate sides facing inwards and then clip into place. It’s a good idea to mark the location of the wells with a permanent marker at this stage to make it easier to see where the wells are later. If running multiple gels it can also be good to mark the top of the longer plate in a way to help distinguish which gels contain what samples.
4. Put the assembly into the clear tank and check that the top part makes contact with the terminals.
5. Fill the space between the gels with 1x SDS PAGE running buffer and fill the tank up to the 2 or 4 gel line depending on how many gels you are running (if concerned about overheating lower weight proteins <25kDa, always fill the tank to the 4 gel line with 1x SDS running buffer). If you only have one gel, use the ‘buffer dam’ on the second side so you get a distinct pool of buffer at the top (an empty gel cassette, tape not removed works well here).
6. Load 5uL of protein ladder standard to one lane (don’t need to boil or add buffer to the ladder). Add the appropriate amount of each of your samples in their respective lanes. Make sure to mark down your loading order! Ensure the gel is asymmetrical as it might get flipped over later during imaging.
7. Run the gel at 200V for 40 minutes or until the dye front reaches the bottom of the gel. Running the gel at a lower voltage for longer may improve the resolution of your gel. Some housings leak so check the gel every now and again and ensure that there is enough buffer between the gels, it may need to be topped up.
8. When the dye has reached the bottom of the gel, stop and disassemble the housing. Used buffer can go down the sink.

#### Visualising SDS PAGE

1. Carefully pry open the plates using a wedge tool or flat head screwdriver. Carefully pry off the gel using gloved fingers or sterile tweezers. It will be reasonably tough but may still tear or break so be careful.
2. Place it in a plastic dish and add enough distilled water to suspend the gel slightly.
3. Next...

1. If you are using a Biorad pre-cast gel with stain free imaging capability move directly to an imager and proceed with the stain-free UV imaging protocol for that machine, transferring the gel carefully to the imaging plate. If you are checking for a protein of interest ensure it contains tryptophan in order to visualize it using Biorad’s stain free technology.
2. If you are not using a stain free gel proceed with coomassie staining by pour off the water into a waste container and adding coomassie blue solution to cover.

1. Put some gladwrap over the top and put on a gentle rocker for half an hour.
2. Pour the stain into a methanol waste bottle and cover with high-destain solution (40% MeOH 10% Acetic Acid) and return, covered, to the rocker for another half an hour.
3. Pour off into the methanol waste and replace with low-destain solution (10% MeOH, 10% Acetic Acid) and rock, covered overnight. The high-destain step may be omitted and done the next morning if there is still a lot of background stain. Don’t worry about a bit of background stain, but less is better as it will enhance the contrast.

4. Gels can be stored in the cold room between two sheets of clear plastic sealed together with tape for later reference or mass-spec band excision. You can annotate the plastic directly but be aware the gel can shift in the plastic over time if not kept flat. If you want to discard the gel, this must be treated as contaminated waste, due to possible acrylamide monomer still present... discard into your chemical waste bag.

Dialysis of protein samples

#### Preparing dialysis tubing.

1. Cut tubing into pieces of convenient length (10-20cm, this will depend on your sample volume).
2. Boil the tubing in 10minutes in a large volume of 2% (w/v) sodium bicarbonate and 1mM EDTA (ph8.0)
3. Rinse the tubing with distilled H20
4. Boil the tubing for 10 minutes in 1mM EDTA (ph8.0)
5. Allow the tubing to cool then store it in 4oC Be sure that the tubing is always submerged

#### Preparing your protein samples for dialysis.

1. Handle with fresh gloves only.
2. Wash the tubing inside and out with sterilized distilled water.
3. Double over one end of the tubing and clip it.
4. Pipette your protein sample in through the open end.
5. Double over the open end and clip that.
6. Submerge your tubing submerged in the solution you wish to dialyse against.
7. Leave overnight at 4oC stirring slowly (magnetic stirrer).

Protocol for heat shock transformation of chemically-competent cells

#### General Notes

This protocol procceds after Digestion & Ligation protocol. This protocol must be done is asceptic condition and will test a users asceptic technique.

1. Remove one or more aliquots (as required) of chemically competent cells of your E.coli strain from
   the -80°C freezer. Thaw the cells e.g. by rubbing them in your hands or put them briefly in a 37°C
   waterbath, but don’t let them stay warm! As soon as they are thawed, put them onto ice.
2. Divide the cells into the appropriate number of 50 µl aliquots in separate Eppi tubes on ice. Add
   your DNA samples to each tube; you can use up to ~10 ul of ligation mixture or plasmid here, but
   note that typically 3 µl of ligation mix or 1 µl of plasmid would be standard.
3. Make sure you include both a positive control and a negative control in the transformation
   experiment. The positive control should be 1 µl of a plasmid with the correct antibiotic resistance
   (same resistance as the plasmid used for the ligation), and should also be a plasmid stock that you
   know is in good condition (based on agarose gel).The negative control is simply no DNA added.
4. Put the cells into a foam ‘floatie’ and put on ice. Ensure at least the bottom half of the tube (approx
   2 cm) is embedded in the ice, don’t just rest them on top of the ice. Allow the cell/DNA mixtures to
   incubate on ice for 15-30 min.
5. Take your esky of ice over to the 42°C waterbath or 42°C heat block. Put the floatie into the
   waterbath. Allow 45 seconds for heat shock. (Plus or minus 10 seconds, this needs to be exact!).
   Then transfer the floatie straight back onto ice (embed into ice, as above, don’t just rest on top).
6. Allow transformation mixtures to sit for 2 min on ice, then add 1 ml sterile LB broth to each tube.
   You can also use more fancy media (e.g. SOC or SOB), but there is not that much difference.
7. Incubate on 37°C shaker for 1 hour. Put the tubes horizontal so they get good shaking action. eg.
   put the tubes laying flat on the shaker platform and masking-tape into place. Make sure the lids
   are tight! You can incubate without shaking, and you can incubate for less time (30 min), but it won’t
   work as well in these cases.
8. Label the LB-antibiotic plates before starting the next bit; you need two plates for each ligation
   condition or plasmid type, since we will plate out two different cell concentrations of each to ensure
   we get countable/pickable numbers of colonies. Double check the plates to ensure you are using the
   correct type of antibiotic(s) for the type of plasmid(s) you are using.
9. Pipette 100 µl of the first cell suspension onto one LB-antibiotic plate (label ‘100 µl’ in addition to
   other info). Sterilise the glass spreader with ethanol and flame (CAUTION! READ THE SOP FIRST!),
   and spread the cells around the plate with the spreader. Do this by pushing the spreader with a backand-forth motion, while turning the plate around in a circular motion. Be careful not to touch the
   spreader on your fingers! Keep spreading for approx 10 seconds.

If the plates are properly dried, you should feel the spreader start to ‘stick’ to the agar, this means
the liquid has been drawn into the agar. If this doesn’t happen after ~20 seconds, stop spreading,
but next time, dry plates for longer! If the plates are incubated with a lot of liquid still on them, you
may not get nice discrete colonies (the cells will swim around in the liquid, making a mess).

10. Spread 100 µl of the remaining samples, each onto a separate, appropriately-labelled plate.
11. Centrifuge all the tubes at ~15,000 rpm for 1 minute in a micro-centrifuge. Pour off most of the
    supernatant into culture waste (being careful not to touch the tubes on the edge of the culture waste
    bottle). Leave a little bit of liquid behind (about one or two drops).
12. Vortex the cells in the remaining liquid for about 10 seconds, until they are not sticking to tube
    anymore, and you have a nice smooth, even, cell suspension.
13. Pipette the cells from the first cell suspension onto the appropriate pre-labelled LB-antibiotic plate
    (label with ‘pellet’ in addition to other info), spread plate as above. Repeat for the remaining samples
14. Incubate all plates at 37°C overnight. Note that for some plasmids and ligations, it may be
    beneficial to instead try room temp for 2-3 days – this lowers the copy number of pUC type plasmids,
    and is useful to allow retrieval of clones that might be toxic to the host.
15. When examining your plates, first check your controls. The positive control should have
    thousands of colonies, perhaps even a confluent lawn of growth, especially on the ‘pellet’ plate. the
    negative control should have no colonies at all. If you don’t see these results with the controls,
    anything you see on your experimental plates is questionable. Common problems and their
    interpretation are summarised in the Table below. Detailed troubleshooting of the different ligation
    controls and their results is described in a different protocol.
“Mix up of labelling” can cause MANY problems – be super careful with your labelling ! (do
this BEFORE starting the hands-on bit of the procedure, and double-check everything)

Problem | Interpretation/solution |
|  |  |
| --- | --- |
| Lots of growth of the negative control (thousands of colonies or lawn) | * Forgot to add antibiotic to the plates * Antibiotic concentration is wrong (too low) * Host bacteria are already resistant to the antibiotic (e.g.   TOP10 has chromosomal streptomycin resistance) * Plates incubated too long (especially with LB-ampicillin) * Severe contamination with an antibiotic-resistant   bacterium (not E.coli) (unlikely!) * Mix up of labelling somewhere – is this actually the   positive control? or one of the experimental tests? |
| Some growth on the negative control (a few colonies) | * Contamination during the procedure, e.g. from one of   the other samples or the pipette etc. This may not be a   ‘deal-breaker’ so long as there are lots more colonies   on your experimental test plates * Mix up of labelling |
| No growth or very little growth on the positive control plate | * The cells are not competent * Used the wrong antibiotic in the agar (check the   sequence of your plasmid to confirm correct resistance) * Used the wrong concentration of antibiotic (too much) * Agar plates are ‘bad’ for some other reason (e.g. added   mercuric chloride instead of sodium chloride!) * Plasmid stock has gone bad (run a gel to check) * Mix up of labelling * Pipetting error (look at the pipette tip to ensure that you   really have 1 µl of plasmid in there!) |

Resting cell assay for cell metabolism

A resting cell suspension contains cells with induced enzymes capable of doing metabolic reactions, but which are suspended in non-growth medium so they can’t make new proteins or cells. Under these non-growth conditions, substrate depletion initially follows zero-order kinetics (linear), allowing simple calculation of apparent specific activities (k) from a straight line fit through the substrate depletion data. Resting cell suspensions are also useful because they allow concentration of cells from a large culture volume into a small volume, thus boosting the reaction rates, and sometimes making reactions measurable that can’t be seen in growing cultures.

#### Protocol

1. Choose a growth medium that you know will give you the enzyme activity of interest.
2. Determine what volume of culture you need to grow to see the activity. If the activity is weak, you may need to grow 500 ml of cells to make a 10 ml resting cell suspension (50-fold concentration factor), but a strong enzyme activity may be visualised in the same volume of culture as resting cell suspension. If the activity of your assay is too high, the cells will consume the substrate too quickly for you to get good quantitative data. If the activity is too low, a linear substrate depletion curve may not be apparent (or you may have to measure depletion over a very long period of time). If you incubate “resting cells” for too long, funny things can happen, for example the cells can uninduce, or other enzymes that you don’t want may switch on.
3. Think about what controls you need. Will the negative control consist of the same strain grown in non-inducing medium? Or will it be an E. coli with vector-only, not the cloned gene of interest? Is there a positive control you can use?
4. Think about replicates. Do you need to grow multiple cultures on the same day? Or should you stagger these over different days? Is it OK to make replicate cell suspensions from the same parent culture...is that still a true ‘replicate’?
5. Think about the need for sterility. Usually, once the culture has grown, we are in the realm of biochemistry rather than microbiology, so it’s OK to use clean but non-sterile glassware and plasticware and solutions etc in a resting cell assay. However, if you want to keep the solutions for a long time, or if the assay runs for a long time, or uses very rich ingredients (for example glucose), you may need to be more careful.
6. Plan the experimental protocol and timing. Make sure you have all the required media, consumables (especially sterile centrifuge tubes), and required solutions. Make sure you are familiar with the way the cultures grow and the analysis methods required before growing up lots of cultures or large volumes of cultures – a quick preliminary experiment is often invaluable to get the feel for an assay before committing a large amount of resources. How long do the cultures take to grow? How long does the assay take to run? Will you need to be in the lab at 3 am on Sunday morning? (if so, maybe rethink when you will inoculate the culture!)
7. Inoculate the cultures from a fresh inoculum (either plate or broth) and grow to mid-log phase. “Mid-log” (or mid-exponential) means different things for different bacteria and different media, but this will typically be around OD600 = 0.5. If in doubt, let the cultures grow to full density, then assume that mid-exponential is about half of that OD600. In some unusual cases, you may need to grow cultures into stationary phase or death phase or even lag phase, if the enzyme you are interested in is induced at those times, but for most common cellular enzymes, you will get best activity when the cells are happy and healthy in mid-log.
8. If you are using wild-type cells which are induced by the growth conditions themselves, skip ahead to the next step. If you are using recombinant cells, where the inducer is artificial (for example IPTG, arabinose, tetracycline), add the inducer now at mid-log phase. Allow to induce for 4- 48 hours depending on the strain, conditions, and enzyme of interest. For E. coli cells and standard induction systems, 4 hr is usually enough, but for mycobacteria or other slow-growers or for unusual enzymes or cold incubations, induction may require a lot longer. Monitoring OD600 during induction is a good idea to get a sense of how induction impacts on growth.
9. Harvest the cultures by centrifugation. Bacteria can be pelleted at quite low g forces and times, e.g. 3000 g for 10 min is fine in most cases. Check the conversion from g to rpm if in doubt – these are usually in the same range (3000 g is about 4000 rpm in a ‘Centaur’ machine 50 ml rotor), but different rotors have different conversion factors. Some cells like mycobacteria may require additives (0.05% Tween-80) to centrifuge properly, if these aren’t already in the medium (and these additives need to be maintained in both wash buffers and the assay buffer). Cells pellet easier at higher salt concentrations; pure water is quite difficult to pellet cells in.
10. WASH THE CELLS! This is important, and often overlooked. If there are significant amounts of rich growth medium (e.g LB) present in your resting cell assay the cells can grow, which will interfere with determining rates of reaction. Washing is done by centrifuging and then resuspending in a buffer like KP (10 mM K2HPO4, pH 7) or MOPS or PBS etc. To resuspend cells, its best to flick or vortex or shake the tube/bottle rather than using a pipette – clumpy cells in particular will stick to tips and be lost. You can shake vigorously, as this won’t hurt most bacteria.
11. More cell washing hints: Don’t forget to add Tween to wash solutions for clumpy cells like mycobacteria (add Tween 80 after to wash solutions after autoclaving). Don’t use plain water as a wash buffer, and ideally use a wash buffer that is the same as the base components in your assay buffer in the later steps. Typically, we would do two wash steps in a convenient volume, for example 20 ml buffer in a 50 ml Falcon tube for culture pellets from a few hundred ml of culture, or 0.5 ml buffer in an Eppi tube for pellets from a few ml of culture.
12. What temperature to use for centrifugation and wash steps? This is a bit controversial. As a starting point, it is best to use a cold centrifuge for both these steps, since this should “stop the cells in their tracks” and minimise any changes their physiology. However, in some cases, the cold treatment may shock the cells, and they may respond better to room temperature centrifugation and storage during setup. In all cases though, avoid them getting hot! (>30 C)
13. After washing, resuspend the cells in a small volume of the same buffer and keep on ice while you are preparing the assays. Though see above, sometimes room temp might be better. By “small volume” here, we mean like 100 x less than the original culture volume. This gives us the greatest flexibility for assay setup. It’s easy to dilute cells further, but annoying to concentrate them. Measure the OD600 of the cell suspension, using an appropriate dilution if the spectro reading is more than 1. Calculate back to figure out the actual OD600 of the suspension.
14. Set up the assay reactions in appropriate types and sizes of tubes. If the substrate is volatile, you will need gas-tight bottles (serum bottles), which are crimp-sealed. Otherwise, you can use McCartney bottles, Eppis or Falcon tubes. If using McCartney bottles or other recycled glassware, wash these with RO water before using, to remove traces of detergent. If the cells need aeration, ensure sufficient headspace (say 75% of sealed bottle volume), or incubate loosely capped. What is your target OD600? (how much activity are you expecting?) – this will determine the volume of the assay, or more specifically, the ratio of the assay volume to the original culture volume. How many samples do you need to take, and what volume are these? (this also impacts the assay volume)
15. Set up a master mix, which contains all the ingredients for your assay (buffer and substrate, maybe cofactors or stabilisers). Aliquot this mix out into your individual tubes/bottles, leaving room to add cells to bring the assays to their full volume. Pre-incubate the master mix at the assay temperature for at least 10 min before starting, then add cells.
16. Immediately take a time zero sample. If this is a liquid sample, centrifuge this immediately in a cold centrifuge, and transfer the supernatant to a fresh tube for immediate assay, or for freezing and later analysis. Label tubes first! Keep in mind that the sampling process itself can interfere with the assay procedure. For example, if you are too slow, the temperature of the incubating shaker won’t be maintained in the cultures during sampling. If you take too much of the initial liquid volume out via sampling, this will change the way volatiles partition in the bottle.
17. Incubate the tubes with shaking at the required temperature. Keep in mind the need for good mixing! For smaller tubes or bottles, it is often better to incubate them horizontally to get good mixing action. This is even more important when assaying volatile substrates, where movement between liquid and gas phases happens. Sample at appropriate intervals – this may be between 1 minute to 1 hour, depending on activity levels.
18. At end of assay, discard cell suspension to autoclave waste, and if using glassware, rinse this out with water straight away (also discard to autoclave waste), so that cells don’t dry onto the glass. If using volatile substrates, you can leave bottles open in fume hood overnight before discarding culture, to allow volatiles to disperse.
19. After doing your assays, calculate the rate of substrate disappearance or product appearance; this will usually be in nanomoles per minute. This can then be converted into an “apparent specific activity” (k) by dividing by the protein content of the assay (mg). We can use previous standard curves to convert OD600 to amount of protein:

* For E.coli grown in LB: protein (g) = 0.21 x wet weight of biomass, and biomass (g/L)= OD600/0.534, therefore protein (g/L) = 0.393 x OD600 or [protein (μg/ml) = 393 x OD600] (Kangwa et al 2015)
* For Pseudomonas putida grown in LB: [protein (μg/ml) = 69 x OD600] (Mai Anh Ly, PhD thesis)
* For Mycobacterium smegmatis grown in MSM-glucose: [protein (μg/ml) = 99 x OD600] (Mai Anh Ly, PhD thesis)
* For Mycobacterium chubuense grown in MSM-acetate: [[protein (μg/ml) = 112 x OD600] (Laura Nolan, Hons thesis)

Cloning DNA into Vector – Digestion and Ligation

#### General Notes

One of, if not the most important protocols of synthetic biology. It’s an essential protocol to memorise and master as most if not all your experiments hinges on the success of this particular method.

A few notes to begin with:

* Quantify the DNA concentration of your chosen plasmid and insert DNA (usually a G-block or a PCR product). This can be done through nanodrop, however keep in mind that the nanodrop DNA concentration result of your plasmid is rough estimate and sometimes incorrect. Therefore you may have to run you’re a digest of your plasmid on gel and estimate relative DNA concentration to the DNA ladder standard.
* Check which restriction buffer is appropriate. For double digests, it’s OK to use a buffer which gives 100% activity for one enzyme, and 75% activity of the other, but lower than this is not good.
* Check the appropriate digest temperature, its usually 37°C, but not always. Search online or on the stock restriction enzyme tube for the proper temperature for digestion.

#### Digestion of Vector and DNA insert

### THE GOLDEN RATIO FOR BETWEEN PLASMID AND DNA INSERT\*

## ~3:1 ratio between Insert DNA Mass: Vector DNA Mass

\*Of course this is influenced by the relative sizes of your vector and insert DNA and you may have to tune this ratio to suite your needs but you want to have AT LEAST a higher ratio of insert DNA to your Vector DNA e.g. ~100 ng of DNA insert: ~25 ng of plasmid = 4:1

#### Plasmid Digestion – Plasmid Master Mix

This particularly useful if you are cloning several genes into the same vector/plasmid.

1. Retrieve 10 x restriction buffer from freezer, thaw completely, and vortex to mix. The same tube of buffer can be used many times, if you are careful with your aseptic technique.
2. Retrieve the plasmid from the freezer, allow to thaw, (e.g. in 37°C waterbath, or rub in your hands, or on bench etc), then put it on ice when it is thawed. It’s not good to leave the plasmid stock at room temp or above for prolonged periods or it may degrade due to traces of nucleases.
3. Retrieve the restriction enzyme(s) from the freezer, put IMMEDIATELY on ice. These are heat-sensitive and you need to look after them. Do not leave them at room temp. Keep on ice while setting up the reaction, then immediately put back in freezer. These don need to be thawed, they are in a glycerol solution which doesn’t freeze at -20°C.
4. Calculate an appropriate mass of Vector DNA for the plasmid digest master mix. For the example below, lets calculate how much volume of plasmid you need to use to get 250 ng – call this ‘X’, and calculate how much water to add to the digest; this is (100 – 10 – 2 – X) – call this ‘Y’.
5. Label your tube(s), then set up the digest(s) by adding the ingredients in the following order. Make sure you use excellent aseptic technique, and change tips every time. It’s OK to set this up at room temp, the reaction tube doesn’t have to be on ice. Mix restriction enzyme buffer + DNA + MQ water mixture by pipetting up and down and centrifuging down before addition of the RE. Enzymes like to be in their correct buffer conditions to function.
   * 10 µl\* of 10x restriction buffer (Should be 10% of your total volume to ensure optimal buffer conditions for digestion).
   * ‘Y’ µl of sterile MQ water
   * ‘X’ µl of plasmid DNA
   * 2 µl of (each) restriction enzyme

   Total Volume: 100 µl
6. Mix by pipetting up and down or by flicking, then tap on bench to get liquid to bottom of tube.
7. Incubate at correct temperature for approx. 2 hours. If it’s a High fidelity RE, 1hr is more than enough. (1 - 4 hr is OK, but overnight digest is too long; this can lead to ‘raggedy ends’ of the plasmid even if it looks OK on a gel; this is due to non-specific nuclease activity).

NOTE: 100 µl seems like a large volume to use, but this helps dilute any impurities in the plasmid prep. Increasing the total volume of the digest and/or reducing the volume of plasmid added often help to improve the quality of a poor digest. If the digested plasmid is for the purpose of ligation, we can use a large volume, since it will be column-purified & concentrated anyway.

#### Digestion of Insert DNA

This will be most likely be G-block from oligonucleotide manufacturer or PCR product with the appropriate restriction enzyme sites at the ends. As said before, you want to have a higher ratio of insert DNA to vector DNA. This digestion can be concurrently performed with the digestion of the plasmid.

1. Calculate an appropriate mass of Insert DNA. For the example below, lets calculate how much volume of Insert you need to use to get 100 ng – call this A, and calculate how much water to add to the digest; this is (15-1.5-1-A) – call this B.
2. Retrieve 10 x restriction buffer from freezer, thaw completely, and vortex to mix. The same tube of buffer can be used many times, if you are careful with your aseptic technique.
3. Retrieve the restriction enzyme(s) from the freezer, put IMMEDIATELY on ice. These are heat-sensitive and you need to look after them. Do not leave them at room temp. Keep on ice while setting up the reaction, then immediately put back in freezer. These don’t need to be thawed, they are in a glycerol solution which doesn’t freeze at -20°C.
4. Retrieve the Insert DNA from the freezer, allow to thaw, (e.g. in 37°C waterbath, or rub in your hands, or on bench etc), then put it on ice when it is thawed. It’s not good to leave the Insert at room temp or above for prolonged periods or it may degrade due to traces of nucleases.
5. Label your tube(s). Set up digest by adding ingredients in the following order. Make sure you use excellent aseptic technique, and change tips every time. It’s OK to set this up at room temp. Mix restriction enzyme buffer + DNA + MQ water mixture by pipetting up and down and centrifuging down before addition of the RE. Enzymes like to be in their correct buffer conditions to function.
   * 1.5 µL\* of 10x restriction buffer (Should be 10% of your total volume to ensure optimal buffer conditions for digestion).
   * ‘B’ µl of sterile MQ water
   * ‘A’ µl of insert DNA
   * 1 µl of (each) restriction enzyme

   Total Volume: 15 µl\*

   \*This volume is quite flexible. Adjust if needed.
6. Incubate at correct temperature for approx. 2 hours. If it’s a High fidelity RE, 1hr is more than enough. (1 - 4 hr is OK, but overnight digest is too long; this can lead to ‘raggedy ends’ of the plasmid even if it looks OK on a gel; this is due to non-specific nuclease activity)
7. Purify both vector and insert DNA using Qiaquick columns (or similar), elute in 15 µl EB.

#### Ligation

1. Retrieve one 10x ligase buffer aliquot from the freezer. This should be a small amount of buffer (e.g. 10 µl) in a small generic tube, not the large tube from the manufacturer with ~1 ml of buffer Ligase buffer (unlike restriction buffer) cannot be repeatedly frozen and thawed, it starts to ‘die’ after even one freeze/thaw cycle.
2. Retrieve the T4 ligase enzyme from the -20°C freezer and put IMMEDIATELY on ice. This reagent is VERY heat sensitive, and must be handled with care.
3. Put your labelled tube(s) on ice, then set up the ligase reaction in this tube on ice. Be careful not to get ice or melted ice in the tube - this is not sterile! Add reagents in this order, change tips each time. Mix restriction enzyme buffer + DNA + MQ water mixture by pipetting up and down and centrifuging down before addition of the RE. Enzymes like to be in their correct buffer conditions to function.
   * 2 µl of 10 x ligase buffer (Should be 10% of your total volume to ensure optimal buffer conditions for digestion)
   * 8 µl purified insert DNA
   * 8 µl purified plasmid DNA
   * 2 µl of T4 DNA ligase enzyme
   Total Volume: 20 µL
4. Mix by flicking briefly then incubate either 4°C overnight or room temp for 1 hour. Return the ligase enzyme immediately to -20°C freezer. Throw out any unused thawed ligase buffer.

If you want to don’t want to deal with column DNA purification you can use:

#### Cloning/ligation protocol using heat-killed digests

This protocol is especially useful for ligations involving very small or large fragments which do not get retained very well during column purifications. It’s also worth considering when you don’t have much DNA in your sample and want to minimise losing it. Note that some restriction enzymes are not heat-killable (e.g. PstI-HF and BamHI) so check this first! (NEB website).

The thermostat on a heat-block isn’t very accurate and usually the block is 5-10°C below the set point. Check the thermometer before you start. Putting the thermometer in an Eppi tube of water in the block will give you the closest idea of what your sample is actually being heated to.

A downside of this protocol is that it doesn’t remove small offcut bits of DNA like column purification does. If for example you are digesting a PCR product that isn’t too small, or a two sites in a plasmid with only a few bases between them, it would be better to column purify these digests to reduce the chance of the small fragment re-ligating back in.

1. While waiting for vector and DNA insert digests, turn on the heating block and set to whichever is the highest heat kill temperature of the two restriction enzymes. When digest is complete, place the tube with DNA into the heating block for 20 min. Turn off heat block! Chill DNA on ice for 5 min.
2. Retrieve one 10x ligase buffer aliquot from the freezer. This should be a small amount of buffer (e.g. 10 µl) in a small generic tube, not the large tube from the manufacturer with ~1 ml of buffer Ligase buffer (unlike restriction buffer) cannot be repeatedly frozen and thawed, it starts to ‘die’ after even one freeze/thaw cycle.
3. Retrieve the T4 ligase enzyme from the -20°C freezer and put IMMEDIATELY on ice. This reagent is VERY heat sensitive, and must be handled with care.
4. Put your labelled tube(s) on ice, then set up the ligase reaction in this tube on ice. Be careful not to get ice or melted ice in the tube - this is not sterile! Add reagents in this order, change tips each time:
   * 2.3 µl of 10 x ligase buffer (Should be 10% of your total volume to ensure optimal buffer conditions for digestion).
   * 10 µl purified insert DNA
   * 10 µl purified plasmid DNA
   * 1 µl of T4 DNA ligase enzyme
   Total Volume: 23.3 µl- Give the tube a quick spin to get all the liquid to the bottom. Ligation can be done at room temperature for 30 min on your bench or overnight in the cold room.

   Proceed with Transformation Protocols!

P450 Expression Culture Method with Chaperone CO-Expression

#### Special Mentions

This protocol was provided to the iGem USYD team by Professor Elizabeth Gillam of
the School of Chemistry and Molecular Biosciences at the University of Queensland. Thanks for sharing this protocol :D!

##### Day 1:

Transform DH5aF'IQTM cells with both the P450 expression plasmid and pGro7, using
either electroporation or ultracompetent cells generated by the Inoue method (see
separate protocol sheet). Alternatively, DH5aF'IQTM pre-transformed with pGro7 can be
stored as a glycerol stock then transformed freshly with the relevant P450 expression
plasmid. Select by growth overnight at 37°C on LBglu/amp/cam plates.

##### Day 2:

1. Put on starter cultures: LBglu/amp/cam of appropriate volume (eg. ~3 ml for 0.5 ml to
   seed 50ml bulk culture) inoculated from frozen transformed glycerol stock or
   PREFERABLY a single colony from a LBglu/amp/cam plate of freshly
   transformed cells. Grow overnight at 37°C, max shaking speed, to saturation.
2. Make up 1L TB base, aliquot and autoclave 50mL TB broths in 500mL flasks in which the bacteria will be cultured. P450 yields are better in
   small scale cultures than in larger batch cultures.

##### Day 3:

1. Add first lot of additives (PREINDUCTION PHASE) to each culture flask.
2. Inoculate these bulk cultures containing TB base and additives for preinduction
   phase as  soon as possible in morning with 1/100 volume of starter cultures ie.
   0.5 ml starter culture  per 50 ml of expression medium.
3. Incubate cultures for five hours at 25°C, shaking at ~ 160-180rpm. This is the time
   required  for the pre-induction phase.
4. At t = 5hrs, add second lot of additives (IPTG, ALA, and ARA) to initaite induction of protein expression. Continue to incubate at 25°C, ~160-180 rpm for a further 43 hours (i.e. harvest at t = 48 hours)

##### Day 5:

1. Harvest cultures at t = 48 hours and prepare spheroplasts as normal (see other protocol sheets).

#### Specific Media

##### LBglu/amp/cam Media - per 1 litre

* 10g bactotryptone
* 5g Yeast Extract
* 10g NaCl
* 0.3 mL/L 10M NaoOH
Autoclaved to sterilise. When cool, add 10ml/l of 20% w/v glucose (filter sterilised). Immediately before use add 1ml/l of 100mg/ml ampicillin and 1 ml/l 20mg/ml
chloramphenicol (from methanolic stock).

##### LBglu/amp/cam agar plates

Prepare as above but add 15g/l bactoagar prior to autoclaving.
When media has cooled to ~ 60°C or less, add glucose, amp and CAM and mix well before
pouring plates.

##### TB (Terrific Broth) BASE MEDIA

Per litre

:

* 12g bactotryptone
* 24g yeast extract
* 2g bactopeptone
* 1ml 1M NaCl
* 40ml 10% glycerol
Make up to 900 ml with milliQ water and autoclave.

##### Notes

Try to make up TB as soon as possible before use and  ALWAYS AUTOCLAVE
IMMEDIATELY AFTER PREPARING BASE MEDIA!  Unautoclaved TB goes off VERY
rapidly! If you absolutely have to store it before sterilising, keep at 4°C for the minimum
time possible and observe carefully before sterilising and using. Keep at RT after sterilising.
Always aliquot base media into the flasks in which you will culture the bacteria prior to
autoclaving. Don't make up a bulk mixture, autoclave, then subaliquot into sterile flasks,
since this increases the chance of introducing contamination

#### ADDITIVES FOR PREINDUCTION PHASE

Immediately before starting induction, add (aseptically, per litre complete TB medium (i.e.
adding to 900ml TB base)):

* 100 ml KPi for TB
* 1ml 1M Thiamine \*
* 250µl trace element solution \*
* 1ml 100mg/ml ampicillin
* 1ml 20mg/ml chloramphenicol

##### NOTE:

Medium with bactotryptone, yeast extract, bactopeptone, glycerol, KPi for TB, thiamine,
trace elements, ampicillin and IPTG is the basic start point for optimising expression - make
modifications on this as appropriate for any given trial.

##### KPi for TB:

* 23.1g KH2PO4
* 125.4g K2HPO4
Make up to 1 litre with milliQ water and autoclave separately. DON'T AUTOCLAVE KPi + TB base media, it will precipate into a crappy mess.

##### Trace element stock:

* 2.7g FeCl3.6H2O
* 0.2g ZnCl2.4H2O
* 0.2g CoCl2.6H2O
* 0.2g Na2MoO4.2H2O
* .1g CaCl2.2H2O
* 0.1g CuCl2
* 0.05g H3BO3
* 10ml conc. HCl
Make up to 100ml with milliQ and autoclave.

These salts preciptate out of solution easily. Therefore needs to directly to medium and mix.

##### Antibiotics & ETC

Ampicillin: 100mg/ml sterile milliQ water
make up immediately before use and filter to sterilise

Chloramphenicol: 20mg/ml in high quality methanol (CAM is very insoluble in water)

1M Thiamine: 337mg / ml sterile milliQ water
make up immediately before use and filter to sterilise

### Notes:

Thiamine, CAM and Amp can all be added to KPi; mixture then added in one step to
medium. e.g. if have a total of 900 ml TB base, autoclave 100 ml of KPi for TB, then filter
sterilise 1ml amp, 1 ml CAM and 1ml thiamine into the KPi, then add 5.15 ml of this to 45ml
TB base). Add trace elements separately (12.5µl per 50 ml culture).
If using previously-opened stocks (eg trace elements) it is safest to filter sterilise an aliquot
again immediately before using. Avoid adding supplements to TB until just before
inoculation with culture, since this medium is rich and will easily grow (other types microbes.

#### ADDITIVES FOR INDUCTION

Following the 5hr preincubation, cultures will be supplemented with (per litre)

* 1ml 1M IPTG (1000X stock: typically 50µl/ 50 ml culture)
* 1ml 500mM d-ALA (1000X stock: typically 50µl/50ml culture)
* 10ml 400 mg/ml arabinose (100X stock: typically 500µl/50 ml culture)

500mM d-ALA: 84 mg/ ml milliQ - make up immediately before use and filter to sterilise
(d-ALA appears to be essential for certain forms so we add for all).

1M IPTG (Isopropylthiogalactoside) 238.2mg / ml sterile milliQ water
make up immediately before use and filter to sterilise

400mg/ml arabinose (ARA)  make up immediately before use and filter to sterilise
Ara may require brief incubation at 37°C or under a hot water tap to dissolve.

These can be mixed together in the appropriate proportions (1:1:10 IPTG: ALA: Ara) and filter
sterilised and added in one lot (e.g. 600µl / 50 ml culture) to each culture flask.

Golden Gate Assembly for Cloning One or More Inserts Into a Vector

#### General Notes

Golden Gate Assembly (GGA) is used for cloning one or more inserts into a vector
by designing overlapping ends which allow the combination of multiple inserts
through the use of a non-palindromic restriction enzyme, BsaI, which is then followed by ligation and transformation.

#### Pre-cutting of inserts and vector

1. Add 1µL 10x Cutsmart (NEB) buffer (1x buffer contains 50 mM Potassium
   Acetate, 20 mM Tris-acetate, 10 mM Magnesium Acetate, 100 µg/ml BSA, pH
   7.9 at 25°C) to a 1.5 mL tube.
2. Add 1µL BsaI-HF (200U)
3. Combine Equimolar amounts of inserts and vector to the tube.
4. Set thermocycler to 37°C and incubate BsaI-HF DNA mixture for 15 minutes

#### Ligation

1. After the BsaI-HF DNA mixture or GGA (Golden Gate Assembled) sample was been incubated, calculate a volume of GGA sample (X) and a volume of MQ water (Y) to make up to a total volume 15µL with:
   * 1µL 5x Cutsmart Buffer
   * 1.5µL 10mM ATP
   * 1µL T4 DNA ligase
   * GGA sample 'X' µL
   * MQ 'Y' µL
   Total volume = 15 µL
2. Run samples on the following thermocycler program with the folowing program parameters:
   1. 37°C 2 minutes
   2. 16°C 2 minutes
   For 25 cycles3. Then 75°C for 15 minutes
   4. Then hold (stasis conditions) for 4°C

Clean up step - This step isn't neccassary but it's purpose is to digest un-ligated free floating inserts. for

1. After running your GGA sample in the PCR conditions above, add 1uL 5x Cutsmart Buffer, 1uL BsaI, and 3uL MQ water (Total Volume is now 20uL!)
2. Set thermocycler to 37°C and incubate BsaI-HF DNA mixture for 15 minutes

3. Transform 3-8 uL of GGA sample to a suitable competent cell line.
4. And we are done... Why are you still here?

Fluorescence Quantification Protocol

There are many ways to go about this, but this is roughly how we did it. If you are using GFP-family protein or a protein with similar excitation and emission wavelengths, I suggest using the iGEM Measurement protocol. But without further adieu.

After screening your fluorescent colonies by PCR and checking correct sequence of the plasmid by Sanger sequencing. Select your purified colonies, and if your vector is inducible, induce them at time (t=0) or just before chuck at the bad boi (plate reader).

*Culture in 5ml LB broth overnight at 37C 200rpm shaking. Ensure you are also culturing a negative Control (Empty/parental vector) and a positive control such as known fluorescent culture expressing a protein with similar excitation and emission spectrums, or the wild-type version of your expressed fluorescent protein.*1. Measure OD of overnight cultures dilute to OD600 0.02 using LB media with appropriate antibiotics and hold on ice (limits growth).
2. IN VERY ASCEPTIC CONDITIONS e.g. A biosafety cabinet; In a 96 well plate, aliquot 100 uL of your chilled cultures into replicates for each of your selected test samples and controls. After aliquoting samples, immediately put back the 96 well plate lid and chill on ice to limit growth and prevent contamination.
3. Set the plate reader, with at a constant temperature and shaking, to measure OD600 and Fluorescence (your specific excitation and emission wavelengths) every 15 minutes for 18-24hrs.
4. Then process your resulting data using your favourite spreadsheet processor! HAVE FUN :D!

Notes: You can typically disregard the first reading as condensation impacts the the OD600 and fluorescence readings at t=0.

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# Team:Sydney Australia/Results

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* Achievements

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Results

[Characterization of PsiD/K/M in vitro](#openModal1)

[Close](#close "Close")

## Aims

In this experiment, we had two aims that we wished to accomplish.* To express psiD, psiK and psiM genes individually in *E.coli*, replicating their known in vivo expression (Fricke, Janis, Blei, Felix, and Hoffmeister, Dirk. “Enzymatic Synthesis of Psilocybin.” Angewandte Chemie International Edition 56.40 (2017): 12352–12355. Web.).
* To characterise the functionality and kinetics of these psi genes in vitro.

## Constructing psi gene Expression Vectors

Our first step successful step in the experiment was cloning of psiD, psiK and psiM genes individually into pET28 vectors (Figure 1). It was important for us to generate stable recombinant pET28 vectors as this would enable us to transform any T7 polymerase containing *E.coli* cell line and induce strong expression of our proteins with an added N terminal 6xHIS tag.

![](https://static.igem.org/mediawiki/2019/9/95/T--Sydney_Australia--PlasmidConstructionDKMResults.png)

Figure 1. Double restriction digestion of psi genes psiD, psiK and psiM in the pET28c vector. Constructs were digested for 2 hours at room temperature alongside an empty pET28 vector and no template control using NEB's HindIII-HF and EcoRI-HF enzymes. Products were loaded onto a 1% Agarose, 40mL TAE gel prestained with Gel Green. The gel was run for 40 minutes at 100V before being imaged by UV transillumination using a BioRad XRS+ Chemidoc and QuantityOne's basic imaging software.

After using our constructs to transform BL21 *E.coli*  cells we started testing our cell lysates for the presence of our proteins after induction. For this, we used SDS-PAGE gel electrophoresis as our diagnostic tool. In our first series of investigations, we encountered a noticeable problem. When we attempted expressing Psi proteins they appeared highly insoluble under our expression conditions (figure 2). To improve the solubility of our protein we began revising and troubleshooting our expression conditions and enlisted assistance from the Macquarie iGEM team (Figure 3) (See more about their help here!( [See more about their help here!](https://2019.igem.org/Team%3ASydney_Australia/Collaborations))

## Achieving soluble Psi protein expression

![](https://static.igem.org/mediawiki/2019/7/75/T--Sydney_Australia--InsolubleProteinExpressionDKMat37Results.png)

Figure 2: SDS PAGE of cell lysates from BL21(DE3) cells containing pET28-psi gene constructs grown the presence or absence of IPTG (1mM). Psi proteins highlighted in red, (A) PsiM, (B) PsiK, (C) PsiD.  Cells were grown to OD600 of ~0.5 at a temperature of 37℃ with shaking (200rpm) before IPTG induction at room temperature. Cultures were then returned to incubate with shaking overnight. Lysates were collected via bead beating. Insoluble lysate fractions were collected by beating cells in 1x SDS sample buffer (4% SDS, 20% glycerol, 0.002% Bromophenol Blue, 5% fresh B-mercaptoethanol, 62.5mM Tris-HCL pH6.8). Soluble fractions were collected by beating cells in 1x TE buffer (10mM Tris, 1mM EDTA pH 8). Gels were run at 200V for 40min. Gels were imaged using BioRad’s stain free UV imaging technology.)

## Insoluble expression at 22°C

![](https://static.igem.org/mediawiki/2019/5/57/T--Sydney_Australia--InsolubleProteinExpressionDKMat22MQResults.png)

Figure 3. SDS PAGE of cell lysates from BL21(DE3) cells containing pET28-psiD gene constructs grown the presence or absence of IPTG (1mM). Cells were grown to OD600 of ~0.5 at temperature of 37℃ with shaking (200rpm) before IPTG induction at room temperature. Cultures were then returned to incubate with shaking overnight at 22℃ . Cells were lysed via sonication (60% amplitude, 6 cycles, 10 seconds on, 10 seconds off) and soluble and soluble fractions were separated by centrifugation (30 minutes, 20,000g) before being run on SDS PAGE)

With the results obtained from Macquarie and our own data we determined that it might be necessary to modify not only our expression conditions but also our system to hopefully increase the soluble expression of our proteins.

We decided to co-transform our pET28 vectors alongside an additional vector (pGro7) encoding the molecular chaperone complex groES-groEL into new BL21 cells. We believed the added chaperone and an incubation temperature around 15C after induction would reduce the stress on our cells and increase the solubility of our proteins. After generating new SDS PAGE gels of our cell lysates we observed evidence of our Psi proteins appearing relative to our uninduced controls in our soluble cell extracts.

![](https://static.igem.org/mediawiki/2019/a/ab/T--Sydney_Australia--SolubleProteinExpressionDKMatResults.png)

Figure 4.SDS PAGE of cell lysates from BL21(DE3)(pGr07) cells containing psi genes present in pET28 vectors in the presence or absence of IPTG (1mM).  Soluble Psi proteins highlighted in red boxes. Cultures were grown to OD600 of ~0.5 at 37 degrees before induction at room temperature. Cultures were then left to incubate at 15C overnight. Cells were lysed by bead beating in 1xTE buffer (10mM Tris, 1mM EDTA, pH 8). Samples were then denatured (3 minutes at 95C in 1XSDS sample buffer: 4% SDS, 20% glycerol, 0.002% Bromophenol Blue, 5% fresh B-mercaptoethanol, 62.5mM Tris-HCL pH6.8) before being run on SDS PAGE. Gel was run at 120V for 60min. Gels were imaged using BioRad’s stain free UV imaging technology.)

## Confirmation of Soluble Psi Proteins

![](https://static.igem.org/mediawiki/2019/2/29/T--Sydney_Australia--ProteinLCMSconfirmssolublePsiKandPsiMResults.png)

Figure 5. SDS PAGE gel containing soluble cell lysate profiles from cultures believed to be expressing soluble PsiK and PsiM was stained with coomassie blue and sent for MS analysis. The bands highlighted in red were cut, destained with 40% acetonitrile/60% 20mM ammonium bicarbonate, dehydrated, rehydrated with 12ng/ul porcine trypsin at 4C for 30min, excess trypsin was removed and 10ul 20mM am.bic was added and incubated overnight at 37C. Peptides masses were determined using MS and samples were identified using a Mascot peptide mass fingerprint search against all si genes.)

We were successful in selecting and identifying two gel spots we believed to be Psi proteins. Mascot returned significant matches with PsiM and PsiK for these bands and although we were unable to identify PsiD, and in the interest of time, we were confident that it was also expressing at our new conditions based on the results for the other two proteins. Using our new expression conditions we grew large cultures of cells containing our constructs and attempted to nickel column purify our Psi proteins from cell lysates. Due to technical errors in the lab, we were only able to recover column purified fractions we believed contained PsiD and PsiK. In the interest of time we decided to test the functionality of these Psi proteins using a qualitative LC/MS in-vitro assay modelled off Frick et al (Figure 6) (Figure 7).

![](https://static.igem.org/mediawiki/2019/3/36/T--Sydney_Australia--psiDChemicalLCMSdataResults.png)

Figure 5: Qualitative LC/MS profile of PsiD in vitro showing conversion of tryptophan to tryptamine. Ni+ Column purified fractions of cell lysate believed to contain PsiD were incubated at room temperature for 2-3 in the presence and absence of L-tryptophan (2.5mM). Ion peaks of interest from in vitro samples, tryptophan (209.0572 m/z) and tryptamine (161.1073 m/z), were identified against known 1mM standards also identified using LC/MS.

![](https://static.igem.org/mediawiki/2019/3/36/T--Sydney_Australia--psiKChemicalLCMSdataResults.png)

Figure 6: Qualitative LC/MS profile of PsiK in vitro showing conversion of 4-hydroxytryptamine to norbeocystin.Ni+ Column purified fractions of cell lysate believed to contain PsiK were incubated at room temperature for 2-3 in the presence and absence of 4-hydroxytryptamine (2.5mM), ATP (5mM), MgCl2 (5mM) and B-mercaptoethanol (5mM). Ion peaks of interest from in vitro samples, tryptophan (209.0572 m/z) and 4-hydroxytryptamine (177.1022 m/z), were identified against known 1mM standards also identified using LC/MS. Norbaeocystin standards could not be obtained.

## Did we meet our aims?

Our LC/MS data yielded exciting results! We were able to confirm catalytic activity for both PsiD and PsiK with PsiD successfully converting L-tryptophan to tryptamine and psiK converting 4 hydroxytryptamine to norbaeocystin in vitro. We noticed the presence of a hydroxylated tryptamine compound in our PsiD assay supplemented with tryptophan. Because we also observed the formation hydroxytryptamine compound in our 1mM standard of tryptamine we suspect that tryptamine may spontaneous oxidize to form this compound and therefore its presence in our in vitro assay is not the result of any conversion by endogenous *E.coli* proteins. Along with our peptide mass fingerprint identification, we made considerable headway in achieving our aims as outlined below.

![](https://static.igem.org/mediawiki/2019/8/8e/T--Sydney_Australia--TablePsiDPsiKPsiMResults.png)

Table 1. Summarised PsiD, PsiK and PsiM Results

## Future Directions?

Going forward with this experiment there are several avenues we wish to explore. We would first want to confirm soluble PsiD expression using LC/MS and peptide mass fingerprint identification. Next we would work towards purifying and quantifying all psi genes once more and use LC/MS again to characterize the functionality of PsiM. We would also like to quantify the amount of Psi protein present in our purified protein fraction. This would enable us to make estimations on the amount of protein present in each cell. Following this, we could then characterize the kinetics of our proteins using HPLC LC/MS. Sampling our in vitro assay containing a known amount of protein during a time course, after the addition of respective substrates, should help us build a kinetic profile for our enzymes. This information would be highly valuable in helping model systems designed using these parts.

[Assembling PsiD, K, and M Gene Cluster for Magi.Coli](#openModal2)

[Close](#close "Close")

## Aim

To take the three genes psiD, psiK and psiM from the psilocybin enzymatic pathway that we had previously expressed using the PK18 vector and clone them into our in-house PUS250 vector which has a tightly regulated cumate promoter system and a chromoprotein allowing easy selection. We built the new construct using Golden Gate assembly and validated our results by Sanger sequencing and LCMS of metabolites.

![](https://static.igem.org/mediawiki/2019/f/f2/T--Sydney_Australia--PUS387spanningpcr.png)

Figure 1: We completed a colony PCR on transformants from Golden gate assembly using primers that spanned the insert. The above gel (1% agarose 1x TAE 40 mM. Tris-acetate and 1 mM EDTA pH 8.5, 100V, 1 hour) shows DNA from the two colonies numbers 14 and 16 amplified with forward primer MVS48 reverse primer CFM23 these were approximately the right size at ~3500bp so we grew up cultures of these colonies and purified plasmid DNA for sequencing.

![](https://static.igem.org/mediawiki/2019/9/9b/T--Sydney_Australia--pUS387ChemicalLCMSdataResultsV2.png)

Figure 2: We completed resting cell metabolism assays on our two PUS387 constructs by growing up cultures of DH5a cells including the PUS387 vectors then transferring them to a Phosphate glucose solution and providing the cells with the intermediate 4-hydroxy tryptamine then allowing them to metabolise at 15C with shaking (150rpm) for 24 hours. The supernatant was then filtered and sent for LCMS analysis. The LCMS results showed peaks at 161.10732 m/z corresponding to Tryptamine the product of PSID activity on endogenous Tryptophan. The Psilocybin precursor Baeocystin was identified as a peak at m/z of 271.08422. This demonstrates the activity of the PsiD, PsiK and PsiM enzymes in the synthesis pathway from 4-hydroxytryptamine.

## Sequencing of pUS387 plasmid

The result of our sequencing shows a large ~400 bp deletion in the psiM part of the construct. This is concerning, as the activity of PsiM must be intact for the production of Baeocystin, which appeared in our LCMS results. This deletion also appears in the sequence of both of our PUS387 constructs which were derived from separate colonies. It seems unlikely that such a large truncation would still result in a functional enzyme and that it could appear in both Golden Gate constructs when we had confirmed the sequence of the inserts was correct. However, diagnostic restriction digest of the PUS387 constructs shows a degraded band of ~500bp which corresponds to the appropriate size of a truncated PSIM protein. There is also the possibility that  *E. coli* contains an endogenous methyltransferase that can catalyse the conversion of Norbaeocystin to Baeocystin. The ability of this protein to still be catalytically active requires further investigation.

![](https://static.igem.org/mediawiki/2019/8/82/T--Sydney_Australia--PUS387digest.png)

Figure 3: Double restriction digestion of pUS250 and pUS387 clones (14 and 16) Constructs were digested for 30 seconds using a 900W microwave alongside a no template control using NEB's HindIII-HF and EcoRI-HF enzymes. Products were loaded onto a 1% Agarose, 40mL TAE gel prestained with Gel Green. The gel was run for 40 minutes at 100V before being imaged by UV transillumination using a BioRad XR+ Chemidoc and QuantityOne's basic imaging software.

## Future Directions

Although we have demonstrated that the enzymes in the pathway are active in the construct we were unable to demonstrate the production of Psilocybin. We postulate that this is due to low levels of activity and insufficient production of Baeocystin to drive the equilibrium of the reaction towards the product. Future work will need to focus on the optimisation of the conditions for protein production. This may involve the optimisation of growing media, conditions and the use of chaperone proteins. One potential optimisation pathway could include the use of error-prone PCR and to screen for and improved protein production phenotype.

### Dual Plasmid Assay

We have cloned both the PUS382 and PUS383 plasmids which contain the gene for psiH into DH5 pGro7 and performed a resting metabolism assay but are yet to receive the LCMS results. This is a key part of our experiment to recreate the entire Psilocybin synthesis pathway in a single organism and brings us closer to our aim of bulk production.

### Considerations for replication of experiments

The unusual sequencing results for the construct will need to be resolved by designing primers to span the putative region of the deletion. Additionally, we need to purify the PSIM protein from our constructs and test its activity with a quantitative assay such as HPLC-MS. This will confirm whether there is a true deletion, some other mutation or an issue with our primer design and will allow us to better characterise the construct. If the deletion is confirmed we will need to rebuild the construct with the correct PSID insert and compare their activities to see if we can improve the efficiency of the construct.

[Expressing and Characterising PsiH Enzymes](#openModal3)

[Close](#close "Close")

## Aim

This part of the project aimed to express and characterise the activity of the PsiH enzyme in vivo before putting it into our single-cell system.

## Plasmid Construction

The first step was to excise the human p450 gene from the pCW plasmid and then re-ligate the plasmid back together. This was achieved, and the plasmid pUS381 was created. From there, we could clone in our wild type PsiH gene, making pUS382, and the codon optimised PsiH gene, making pUS383. The maps of these plasmids can be seen in Figure 1, and restriction digests can be seen in figure 2.

![](https://static.igem.org/mediawiki/2019/4/49/T--Sydney_Australia--allplasmidmapsPsiHResults.png)

Figure 1. Plasmid maps of pCW, pUS381, 382 and 383. (a) PCW-CYP2A6 is the parental plamid of the other plasmid constructs. It contains a human p450 (in red) and cytochrome p450 reductase (in brown). (b) pUS381 is a PCW-CYP2A6 plasmid derivative lacking the human p450 gene but still containing the p450 reductase, therefore it was used as the base plasmid from which pUS382 and pUS383 were constructed. (c) pUS382 contains the wild-type psiH from *Psilocybe cubensis* (in purple), while (d) pUS383 contains the IDT codon-optimised psiH (in purple).

![](https://static.igem.org/mediawiki/2019/2/23/T--Sydney_Australia--pUS381-2-3PCR.jpg)

Figure 2. PCR of pUS381, pUS381 and pUS383. Spanning PCR products in the left four lanes show psiH insert in pUS382 and pUS383. Junction PCRs used primers that bound internally to psiH genes. The middle four lanes show products of PCR with primers specific to wild type psiH gene. The right four lanes show products of PCR with primers specific to codon optimised psiH. ntc - no template control.

After constructing each plasmid (pUS381, pUS382 and pUS383), they were sent off for Sanger sequencing. It was confirmed that pUS381 and pUS382 were correct. However, we found out that in pUS383, there was a single nucleotide deletion at the start of the codon optimised PsiH gene. This results in a frame-shift mutation and a truncation of the protein. Unfortunately, this sequencing data only came back after we had performed all of the assays, and we did not have time to re-do them.

## Protein Expression of psiH

To confirm that our protein could be expressed in *E. coli*, we set up cultures of cells containing the plasmids pUS381, pUS382, and pUS383. Each plasmid had two cultures – one that was induced with IPTG and one that was uninduced. The insoluble fraction from these proteins was run on an SDS PAGE gel and imaged with stain-free imaging.

![](https://static.igem.org/mediawiki/2019/e/e3/T--Sydney_Australia--InsolubleProteinExpressionpsiHResults.png)

Figure 3.SDS PAGE gel showing insoluble fraction of cell lysate from induced and uninduced cultures of pUS381, pUS382 and pUS383.  Cells were grown in TB for 43 hours after induction with IPTG. Insoluble fraction was collected by lysing cells at 95°C for 5 minutes in SDS-containing loading buffer, then supernatant was loaded onto precast 8-16% PAGE gels. Gel was run at 120V for 70 minutes and imaged with stain-free imaging. U = uninduced. I = induced. Red boxes show probable PsiH protein.

The SDS PAGE gel clearly shows bands (highlighted with red boxes) that appear in the cell lysate from induced cultures containing PsiH – pUS382 and pUS383. They do not appear in the cell lysates of uninduced cells, or cells that had the pUS381 plasmid with no PsiH. These bands are therefore probably PsiH. PsiH is a 57kDA protein, but because it is a membrane-bound protein, it can appear on gels at a slightly different size. This explains why the PsiH bands on this gel appear to be approximately 50kDa.

As mentioned earlier, the pUS383 construct had a single nucleotide deletion, resulting in a frameshift mutation. This frameshift results in a truncation and brings the molecular weight of the protein down to 52kDa. It can be seen that the PsiH band in the pUS383 induced fraction is slightly lower on the gel than the band in the pUS382 fraction

![](https://static.igem.org/mediawiki/2019/6/67/T--Sydney_Australia--SolubleProteinExpressionPsiHforLCMSResults.png)

Figure 4. SDS PAGE gel sent for Mass Spec analysis. The insoluble fraction of induced cell culture lysates were run on a gel, then stained with Coomassie blue. The bands suspected to be PsiH were cut out, destained with 40% acetonitirile/60% 20mM ammonium bicarbonate, dehydrated, re-hydrated with 12ng/ul porcine trypsin at 4C for 30min, excess trypsin was removed and 10ul 20mM am.bic was added and incubated overnight at 37C. Peptides masses were determined using MS and samples were identified using a Mascot peptide mass fingerprint search against all psi genes.

Another SDS PAGE gel of the same samples was run and stained with Coomassie blue (shown in Figure 4), to be sent off Mass Spectrometry analysis of the protein. The Coomassie staining makes it harder to see the PsiH proteins. However, it does show other relevant proteins. As indicated in Figure 4, the 70kDa cytochrome p450 reductase partner protein, which is also under IPTG inducible control, can be seen in the induced cell lysates. There is also a very large band which is likely to be the 60kDa chaperone proteins, groES, that we are using to help with PsiH expression.

Despite being able to see bands on the gel, the Mass Spectrometry results were unable to identify PsiH. This does not mean that PsiH was not present - rather, it means that PsiH was not the most abundant protein in the portion of gel that was analysed.

## Testing PsiH enzyme function - Indole Assay

![](https://static.igem.org/mediawiki/2019/8/85/T--Sydney_Australia--psiHindoleAssayphotosResults.png)

Figure 5. Cultures of induced cells after indole assay had been performed. From left to right: pCW, pUS381, puS382 and pUS383

After seeing that our PsiH proteins were being expressed we wanted to test their functionality, so we decided to do an indole assay. Many p450 monooxygenases can oxidize indole to make coloured compounds such as indigo. *E. coli* cells contain endogenous indole, so when grown in the right conditions, cells containing functional p450 monooxygenases can turn blue. This assay takes around 5 days and has many complicated components. We used a positive control, the pCW plasmid containing a human p450 monooxygenase which is known to turn indole to indigo so that we could tell if we had performed the assay correctly. We also used a negative control, the pUS381 plasmid that had no p450 monooxygenase. For each plasmid type, we grew up two cultures, and then induced one and left the other uninduced so we could compare the colours. The results of this experiment show that the induced culture of cells containing pCW turned blue, but none of the other induced cultures changed colour (as shown in figure 5). We did not expect to see a colour change in the pUS381 culture, as it contains no p450 enzyme. Retrospectively, it was also impossible that there would have been a colour change in the pUS383 culture either, because of the frameshift mutation in the codon optimised PsiH in this plasmid. Therefore, this assay only gives us information about the wt PsiH gene in our pUS382 plasmids

This assay could reveal one of two things about that wt PsiH gene - either it was expressing but was not functional, or was functional but could not oxidise indole or is doing so in such a small amount that it was impossible to see. In order to determine which of these conclusions is correct, we needed to perform another assay.

## Testing PsiH enzyme function - Tryptamine assay

This assay was performed to test the ability of PsiH to catalyse the reaction that it performs in the psilocybin biosynthesis pathway, the oxidation of tryptamine to 4-hydroxytryptamine.

![](https://static.igem.org/mediawiki/2019/b/b1/T--Sydney_Australia--psiHpUS382pUS383ChemicalLCMSdataResults.png)

Figure 6. LCMS results analysing compounds in media of resting cell assay.  Cells containing pUS381, pUS382 and pUS383 were grown up as per indole assay and induced with IPTG. 8 hours after induction, cells were transferred to phosphate buffer containing 2% glucose, and supplied with 1mM tryptamine dissolved in DMSO. Samples were collected 24 hours after addition of tryptamine, pelleted, and substrate sent off for LCMS analysis. 1mM samples of tryptamine standard and 4-hydroxytryptamine were also analysed with LCMS for comparison.

The LCMS results were difficult to analyse. The reason for this was that a substance of the same molecular mass and composition as 4-hydroxytryptamine was found in our ‘pure’ tryptamine standard. We also saw this compound in our negative control (cultures with pUS381) and in our culture with pUS383, which we now know has a mutated psiH. However, we did not see any of this compound in our culture expressing the correct PsiH. This compound may have been 4-hydroxytryptamine, but may also have been 5 or 6-hyroxytryptamine - LCMS is unable to distinguish between them. These results are unexpected and contrary. Firstly, they show that our PsiH gene was probably non-functional. Secondly, they may show that tryptamine can auto-oxidise to some compound that is the same or very similar to 4-hydroxytryptamine. This can potentially confuse any attempted analysis of PsiH function with this assay.

## Conclusions

Ultimately, while we were able to induce *E. coli* cells to express PsiH, it was not found to be functional on either indole or its substrate, tryptamine. This result is not surprising, as we thought that PsiH would have to be codon optimised or harmonised to work in E. coli. Unfortunately, we were unable to test the codon optimised PsiH function in *E. coli* due to a frameshift mutation in the gene that drastically changed the coding sequence.

## Future directions

Future directions of PsiH should be focused on further adapting the sequence of the gene for better expression in E. coli. This could be achieved by modifying the N-terminal of the gene and trying different harmonisation and optimisation methods. Additionally, different p450 reductase partner enzymes could be tested along with PsiH, such as fungal variants.

[Creation of Two-Plasmid System](#openModal4)

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#### Aim

Express PsiD, PsiK, PsiM and PsiH in DH5-alpha and assess enzyme function using LC/MS.

![](https://static.igem.org/mediawiki/2019/1/17/T--Sydney_Australia--DualPlasmidSystemResults.png)

Figure 1: Colonies on LB-Km-Cm-Carb plates for PUS387-PUS382 cells.

We were able to successfully transform our chassis with our two plasmids, pUS387 and pUS382. With this transformation, we now had our first iteration of Magi.coli! - An organism with all wild type psi genes. This was as far as we got with our two plasmid system. We set up metabolic resting cell assays using cultures containing these constructs, however we have yet to gather LC/MS data for this assay.

While we have not achieved our ultimate aim with this experiment so far, we are happy with our progress :D.

## Future directions

The most immediate direction we can take with this experiment would be performing LC/MS on our resting cell assays in the presence of excess tryptophan and determine how functional the system is.

[Culturing the Magi.coli Chassis: Lab Vs Fermenter](#openModal8)

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#### Aim

The aim of this experiment was to investigate and compare the growth of our chassis organism in two separate systems:

* Fermenter: The culture was grown in batch mode using a BIOSTAT A fermenter (Sartorius). The fermenter had two six-bladed Rushton impellers, a stirrer speed of 750 rpm was used, the airflow rate was 1 vvm (1.5 L/min). A 1.5 L working volume was used, foaming was controlled via the addition of a 2% (v/v) solution of Antifoam A (Sigma) using an automated foam probe. The temperature was maintained at 37 ℃. Temperature and pH were logged using the in-built data-logging software at a rate of one sample per minute (Figure 1).
* Lab culture: The culture was grown at 37oC with shaking at 200rpm in a 2L conical flask. A 1.5L culture volume was used.

![](https://static.igem.org/mediawiki/2019/4/40/T--Sydney_Australia--ShakingIncubatorVsFermentorFig1Results.png)

Figure 1. Dr Dale McClure's Fermentor. Ain't it a beauty? A heavy duty beauty but still a beauty.

![](https://static.igem.org/mediawiki/2019/0/0d/T--Sydney_Australia--ShakingIncubatorVsFermentirResults.png)

Figure 2. Figure 2. 2L fermenter vs 2L conical flask culture growth. 1.5L of TB broth supplemented with phosphate buffer, trace metal elements and thiamine in each system were inoculated with 100mL of DH5-alpha cells grown overnight in glucose supplemented LB. OD600 was sampled periodically and used to determine density (cells/L)

The results of this exercise demonstrated that the fermentation system is vastly superior in cultivating the growth of our chassis in our media of choice. The number of cells per litre of culture grown was 7.43 times higher in the fermenter than the lab culture over 7 hours. Growth in the fermented culture was so effective the culture had exhausted its nutrient supply by hour 6 resulting in a decline steady decrease in cell density.

Going forward we would like to optimize our Magi.coli for expression at these conditions to take advantage of this stark difference in growth.

[Codon Harmonisation Experiments with Vivid](#openModal5)

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## Aim

The aim of this experiment was to determine which type of codon harmonisation/ optimisation (rank-order method, absolute frequency method, relative frequency method and IDT codon optimisation) would improve the expression of the VVD protein compared to the wild-type codon usage. CH1 was harmonised by rank order, CH2 by nearest frequency, CH3 by relative proportion and CH4 used the IPT codon optimisation tool available here. The harmonisation method descriptions can be found under the documentation for software. The results of this experiment would then be used to codon harmonise our psi genes to improve their expression.

![](https://static.igem.org/mediawiki/2019/7/7b/T--Sydney_Australia--vividharmonisationgel.png)

Figure 1. Spanning PCR of the codon harmonised/optimised VIVID inserts in the pk18 vector showing band of correct size at ~624bp. Agarose gel electrophoresis was conducted on 1% agarose gel in 1X TAE for 50 min at 100 V.

![](https://static.igem.org/mediawiki/2019/0/02/T--Sydney_Australia--VIVIDmutants.png)

Figure 2. Wild type, CH1, CH2, CH3, and CH4 compared to M130I under long-wave UV (~350nm) on charcoal LB agar demonstrating fluorescence of all the Vivid protein types.

![](https://static.igem.org/mediawiki/2019/8/8e/T--Sydney_Australia--HarmonisationsOnlygraph.png)

Figure 3. We conducted a fluorescence assay using a TECAN Spark microplate reader Escherichia coli TOP10 cultures containing WT, CH1, CH2, CH3, and CH4 in pK18. As shown the codon harmonisation and optimisation did not improve fluorescence. The wild type vivid remains significantly (p<0.01 post hoc anova with Bon Ferrony test) more fluorescent than the variants we created.

#### Considerations for Replication

Through this experiment, we have concluded that codon harmonisation and optimisation may not be the most efficient method for improving the activity of these fluorescent proteins. We had significantly more success with our error-prone PCR improvement (see Error-Prone PCR of Vivid). Codon harmonisation and optimisation are essential for other expression applications in particular where the speed of translation may need to be limited to ensure appropriate protein folding.

#### Future Directions

We experienced some difficulty in maintaining the stability of the pk18 VIVID constructs in TOP10 cells. This may be since pk18 is a high copy number plasmid with constitutive expression and may have caused stress to the TOP10 cells. Stability may be improved using an Inducible rather than constitutive vector.

Literally 3 months of painful work with failures left and right...

[Error-Prone PCR of Vivid](#openModal6)

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## Aim

The aim of this experiment was to use error-prone PCR to generate a more fluorescence version of the weakly fluorescent Vivid protein from the fungi, *Neurospora crassa*.

[More on the background of this part](https://2019.igem.org/Team%3ASydney_Australia/Improve)

## Error prone PCR

Error prone PCR was performed on the weakly fluorescent codon optimised vivid gene. The PCR products, when ligated in plasmids and transformed into *E. coli*, yielded 12 colonies that were brighter than the rest (Figure 1).

![](https://static.igem.org/mediawiki/2019/f/fe/T--Sydney_Australia--VVD-met301Figure1Experiments.png)

Figure 1: Results from initial ligation and transformation of error-prone PCR products into TOP10 cells. Extra bright colonies are indicated with yellow arrows.

The sequencing results of the vivid genes from these bright colonies revealed that there was a common mutation in the majority of these genes (see Figure 2). This mutation modified the methionine at residue 130 to either isoleucine (found in 5 of the mutants), a threonine (found in three of the mutants), or a leucine residue (found in one of the mutants).

![](https://static.igem.org/mediawiki/2019/6/64/T--Sydney_Australia--VVD-met301Figure2Experiments.png)

Figure 2: Sequence alignment of error-prone PCR mutants with original VVD gene, with the region of common M130 mutations circled.

When a G-Block of the vivid gene containing a Methionine to Isoleucine change at residue 130 was cloned into *E. coli* and compared to vivid genes without the mutation, it was seen to be much brighter (Figure 3). This confirmed that it was the mutation away from methionine at residue 130 that improved the fluorescence of the protein.

![](https://static.igem.org/mediawiki/2019/0/02/T--Sydney_Australia--VIVIDmutants.png)

Figure 3: The Met130Ile vivid mutant compared to the weakly fluorescence wild type vivid and 4 codon harmonised vivid variants (CH1, CH2, CH3 and CH4) under long wave UV light.

A fluorescence assay was performed that compared the original codon harmonised fluorescence Vivid, the wild type fluorescent Vivid, an empty plasmid, and the new brighter Met130Ile Vivid protein (see Figure 4). This shows the extent to which the single amino acid mutation increased the fluorescence of the protein.

![](https://static.igem.org/mediawiki/2019/2/2f/T--Sydney_Australia--M130vsWTvsCH4Experiments.png)

Figure 4: Graph of TECAN fluorescence data from the VVD-Met301 g-block cells and the original VVD\_wt and VVD\_CH4 cells.

Lastly, the excitation and emission spectra of the new Met130I Vivid protein was measured (figure 5). This spectrum is remarkably similar to the spectra of CFP, meaning that our fluoroprotein can be used as a substitute for this fluoroprotein.
![](https://static.igem.org/mediawiki/2019/f/f0/T--Sydney_Australia--ExcitationsEmissionsVVDResults.png)

Figure 5: Excitation and Emission spectra of the Met130Ile Vivid protein.

## Conclusion

Error prone PCR was used to create a more fluorescent version of the fungal protein, Vivid.

## Future directions

Future work should focus on characterising the protein so that its usefulness can be compared to other fluoroproteins can be established.