iGEM 2025 — Team uppsalauniversity

{% extends "layout.html" %} {% block title %}Notebook{% endblock %} {% block
lead %}This serves as a chronological record of our team's progress throughout
the season. It contains weekly summaries of our activities, experiments, discussions, and
decisions.{% endblock %} {% block page\_content %}

[Week 25](#week25)
[Week 26](#week26)
[Week 27](#week27)
[Week 28](#week28)
[Week 29](#week29)
[Week 30](#week30)
[Week 31](#week31)

#### Week 25

##### June 9 - June 13

---

We honed the project plan for this year's iGEM project and contacted
researchers/professors (Petroutsos lab) to get feedback and help with
materials for our project.

Strains such as starch-filled and WT with and
without cell walls and promoters such as salt-induced and heat shock were
also chosen after feedback. Practical/Industrial applications for our
project were discussed.

We also had some lectures on BioBricks, biofuels
and biosensors (Forster, Danielson). There was also a course on waste
management where we were taught the correct disposal of chemical,
biological and general lab waste.

We started planning the Swedish iGEM
Conference which we are hosting this year. We will have teams from
Stockholm, Lund and Gothenburg coming to Uppsala to discuss ideas and connect.

The team was divided into subgroups responsible for the Wiki,
Swedish Meet, Team Building, Co-operations, PR, Sponsorship,
Documentation, Graphical Design, Video, and Human Practice. Lab groups
were made for Induction, Flocculation Genes, Pair Flocculation, Modelling,
and Assay Prep which will work in parallel during the summer.

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#### Week 26

##### June 16 - June 19

---

A shorter week due to Midsommar

This week we designed the Z17/18 cassettes for transformation into Chlamydomonas
including codon optimization and intron insertion. After which we designed primers for
Gibson assembly into the pLM005 plasmid, and validated them in Benching.

We were given an in-person tour of the Petroutsos lab to familiarize the team with
available equipment and space and claimed Course Lab 5 for future cultivation work.
After seeing the resources available for us we developed written protocols to familiarise
ourselves with working with Chlamydomonas (sedimentation, flocculation, growth curves).

Work was also conducted in E. coli, as we began experimenting with transforming Z17 and
Z18 into E. coli, including low-volume DNA transformations in DH5α strains. We purified our
pLM005 plasmid from the E. coli colonies we received from the Petroutsos Lab.

The Human Practices group continued their research and documentation tasks and performed
further outreach to companies. The Team Building group finalized the date and scope for
the Swedish Meet and the Social Media group drafted a production plan and script for the
promotional video, designed multiple logo versions and researched vendors for team apparel.

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#### Week 27

##### June 23 - June 27

---

This week we focused on preparing key genetic parts, running transformations,
and advancing our algae experiments. We completed multiple E. coli transformations with Z17/Z18,
tested transformation efficiency, and made more glycerol stock. We also ran digests and gels
(including pLM005), performed minipreps, and prepared plasmids for electroporation into
Chlamydomonas. We finalised the design of the FLO1 plasmid, which needed to be split into
three parts for ordering; thereby requiring us to design it for reassembly using Gibson assembly,
designed and ordered primers from IDT, and had a meeting to refine the assembly strategy.

On the algae side, we started to collect data to plot growth curves for our Chlamydomonas
strains, started additional cultures for nitrate and cell-wall-less strains, and ran
experiments to attempt to quantify the sedimentation rate.

Additional work included planning Human Practices, writing a project abstract for
the Maastricht journal, designing PCR primers for the GP1 gene, and continuing our
team's outreach materials (photos, Instagram presentation template, logo work).

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#### Week 28

##### June 30 - July 4

---

##### **What we did this week:**

* Refreshed algal cultures and prepared them for electroporation
* Identified contamination in TAP media and planned improved storage practices
* Took final measurements for the growth curve experiment and plotted results in RStudio
* Prepared and autoclaved fresh TAP-agar, poured plates for Chlamydomonas
* Performed multiple electroporations of CC-125, refining the protocol with 1 mm cuvettes in EBC buffer
* Plated transformed Chlamydomonas cultures onto agar plates
* Maintained overnight cultures of pLM005 for plasmid work
* Completed digestions of pLM005 with XbaI, BamHI, and HpaI, with successful agarose gel analysis and fragment purification
* Purified pLM005 and mRFP1 plasmids
* Ran SDS-PAGE gels to assess Z17 purification
* Conducted colony PCR (cPCR) and analysed products via gel electrophoresis
* Researched the Fus1-Mar1 receptor pair as an alternative flocculation strategy
* Held gene design and Human Practices meetings to align subteam goals
* Sent our abstract to Maastricht for submission
* Shared lab progress by posting presentations on Instagram to engage the public and document our journey

##### **Reflections**

This week was a true exercise in problem-solving, persistence,
and teamwork. We faced contamination setbacks but adapted quickly,
used repetition to master electroporation, and strengthened our molecular
workflows with careful documentation and analysis. Our meetings helped keep
everyone aligned, and our social media updates reminded us of the importance of
sharing science openly. Overall, we kept momentum despite challenges, demonstrating
the resilience and collaboration at the heart of our iGEM project.

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#### Week 29

##### July 7 - July 11

---

##### Weekly Progress

* **Z17-GFP Protein Expression:**
  + Performed IPTG-induced expression in BL21 cells.
  + Several SDS-PAGE gels were run, and no strong Z17 band was detected initially.
  + Explored urea-based extraction for improved solubility.
  + Conducted mini-preps and digestions of Z17-GFP and Z18 plasmids in
    various E. coli strains (DH5α, JM109).
* **Sedimentation & Flocculation Assays:**
  + Carried out tests on wild-type and starch-producing strains.
  + Began data processing and analysis.
  + Investigated the effectiveness of cell counting for quantification.
* **PCR & Cloning:**
  + Designed and ordered primers for GP1 and the expression cassette.
  + Troubleshot PCR with longer extension times; positive controls worked.
  + Ran agarose gels for PCR analysis.
  + Prepared for colony PCR for GP1.
* **Strain Work & Transformation:**
  + Initiated new BL21 cultures for protein comparison.
  + Electroporation plans began for membrane-deficient Chlamydomonas strains.
  + Wrote draft electroporation protocol and started corresponding salt stress experiments.
* **Human Practices & Documentation**
  + Planned Human Practices activities.
  + Began finalising some protocols for addition to the wiki
  + Published presentations and social media stories, including a new
    Facebook profile picture and poster draft.
* **Media & Protocols**
  + Prepared fresh TAP media and agar for Chlamydomonas cultures.
  + Continued adjusting lab protocols and started searching for improved membrane-strand protocols.
* **Research & Investigations**
  + Investigated the salt promoter (optimal concentration for stress experiments).
  + Researched the unknown anchor gene and reviewed FLO1 assembly protocols.
* **Team & Organisational Updates**
  + Held an association meeting and elected new board members.
  + Learned to use the microscope at EBC, capturing images for documentation

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#### Week 30

##### July 14 - July 18

---

##### Weekly Progress

* **Z17/Z18 Protein Expression & Analysis:**
  + Performed IPTG induction of Z17-GFP in *E. coli* BL21.
  + Conducted multiple SDS-PAGE gels on induced and uninduced
    Z17/Z18 samples, including urea-based extractions to
    improve solubility.
  + Began troubleshooting protease digestion and TEV cleavage of Z17,
    including plans for size-exclusion chromatography.
  + Prepared glycerol stocks for Z17 and Z18 strains and
    performed urea extractions under various conditions.
* **PCR, Cloning & Assembly:**
  + Ran multiple PCRs, including FLO1, Z17 Gibson cassette,
    and *Chlamydomonas* lysate templates optimized for high GC content.
  + Successfully visualized PCR bands and continued troubleshooting where necessary.
  + Performed digestion and ligation of Z17-GFP/Z18 fragments (XbaI, BamHI),
    with ongoing troubleshooting of pLM005 ligation using different T4 ligase kits.
  + Ran agarose gels for verification and troubleshooting of PCR, digestion,
    and Gibson assembly results.
* **Transformation & Strain Work:**
  + Performed multiple electroporations of wild-type and
    membrane-deficient *Chlamydomonas reinhardtii* at different
    voltages, followed by plating and recovery.
  + Refreshed *Chlamydomonas* cultures in preparation for
    electroporation and subsequent experiments.
  + Transformed Z18 plasmids into *E. coli* and monitored
    overnight cultures and restreaks.
  + Prepared cultures for protein overexpression and
    performed mini-preps on transformed plasmids.
* **Sedimentation & Salt Tolerance Assays:**
  + Continued salt tolerance experiments, including day 5 measurements.
  + Analyzed flocculation and sedimentation results, preparing statistical
    summaries for wiki documentation.
* **Human Practices & Documentation**
  + Drafted plans for future Human Practices activities.
  + Continued extensive wiki work, including uploading protocols,
    building protocol pages, and updating team member contributions.
  + Merged updates into the main branch and prepared summary documents.
  + Created social media content, including Instagram posts, and prepared
    poster drafts.
* **Media & Protocols**
  + Prepared and refreshed TAP media and agar plates for
    *Chlamydomonas* work.
  + Documented and updated electroporation and transformation protocols.
  + Started developing and optimizing new membrane-deficient strain
    electroporation protocols.
  + Began planning for TEV cleavage workflows and protocol integration into
    downstream analysis pipelines.
* **Research & Investigations**
  + Investigated unknown components related to the FLO1 assembly.
  + Continued exploring optimal salt promoter concentrations for stress
    response experiments.
  + Reviewed and optimized PCR strategies, including high GC content templates
    and primer design considerations.
* **Team & Organisational Updates**
  + Worked on team coordination, teambuilding, and planning for
    future outreach activities.
  + Captured microscopy images of *Chlamydomonas* for
    documentation and analysis.
  + Focused on troubleshooting key steps across cloning, protein expression,
    and transformation workflows to improve reproducibility and yield.

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#### Week 31

##### July 21 - July 25

---

##### Weekly Progress

* **Z17/Z18 Protein Expression & Analysis:**
  + Performed IPTG induction of Z17-GFP and Z18 from overnight cultures and
    recent transformations.
  + Prepared buffers and carried out purification of Z17 protein using Ni2+
    affinity chromatography with two different protocols.
  + Analyzed protein purification results via SDS-PAGE, with further gels
    planned for follow-up validation.
  + Continued exploring protein refolding from urea extractions to
    improve yield and solubility.
* **PCR, Cloning & Assembly:**
  + Ran PCR for Z18 cassette and GP1 constructs, followed by gel electrophoresis
    for verification.
  + Digested pLM005 and Z17-GFP plasmids using multiple restriction enzymes
    (HpaI, BamHI, XbaI) and performed ligation reactions with Z18 inserts.
  + Conducted Gibson assembly for FLO1 constructs with the salt promoter and
    the “China” sequence, followed by transformation.
  + Troubleshot ligation issues involving Z18 and pET plasmids, including
    control reactions and alternative strategies.
* **Transformation & Strain Work:**
  + Completed transformations with ligation and Gibson products.
  + Restreaked all four *Chlamydomonas* strains for continued
    use in downstream experiments.
  + Maintained growth curves across multiple strains and performed cell
    counts to support biomass and flocculation analyses.
* **Sedimentation & Salt Tolerance Assays:**
  + Planned and prepared for large-scale sedimentation and flocculation
    experiments with *Chlamydomonas*.
  + Conducted first-round assays and repeated them with improved timelapse
    imaging to track sedimentation dynamics.
  + Continued refining experimental conditions for quantitative
    comparisons between strains.
* **Human Practices & Documentation**
  + Continued filming and documentation of experiments for
    communication and Human Practices outputs.
  + Took updated team and lab photos for wiki and outreach purposes.
  + Maintained detailed records of cloning, transformation, and purification protocols for
    future reproducibility.
* **Media & Protocols**
  + Prepared necessary buffers and reagents for protein purification
    and column chromatography.
  + Optimized purification workflows and planned improvements based on early results.
  + Gel-purified digested plasmids to improve downstream assembly and ligation efficiency.
* **Research & Investigations**
  + Investigated protein solubility challenges by testing alternative purification
    strategies and refolding approaches.
  + Evaluated performance of Gibson assembly and ligation approaches, troubleshooting based on
    gel and transformation outcomes.
  + Collected and analyzed growth data for different Chlamydomonas strains to better
    understand biomass dynamics in flocculation assays.
* **Team & Organisational Updates**
  + Coordinated planning for upcoming large-scale experiments and
    data collection phases.
  + Captured new laboratory media (photos and videos) for future
    presentations, wiki content, and outreach.
  + Continued collaborative troubleshooting sessions to resolve ongoing
    cloning and purification challenges.

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{% endblock %}

{% extends "layout.html" %}
{% block title %}Engineering{% endblock %}
{% block lead %}The engineering capabilities of our team are represented by how we dealt with issues attempting to amplify certain genes by PCR.{% endblock %}
{% block page\_content %}

[PCR Troubleshooting](#PCR)
[Primers Used](#primers)
[References](#refs)

### PCR troubleshoot

We planned to have several of the parts needed for the project, such as the GP1, FUS1, MAR1 to be amplified from *C.reinhardtii* genomic DNA, however that turned out to be more difficult than expected. While cloning genes through PCR is a rather straightforward method, the high CG content and highly repetitive genome regions provide a challenge when cloning *C.reinhardtii* genes.Our first approach was to adapt a cPCR protocol from Nouemssi et al. (2020)[[1]](ref1) The cells were boiled for 10 min in EDTA buffer, and the lysates were then used in the PCR. The PCR was done using [Taq DNA Polymerase](https://www.thermofisher.com/order/catalog/product/EP0401)
(ThermoFisher Scientific) kit, following the manufacturers instructions. The PCR targeted the coding sequence for GP1 and used the primer pair GP1-F1/GP1-R1. The samples were then loaded on a 1% agarose gel, in order to visualize the results. We however did not see any product on the gel.

To rule-out the possibility, that the reagents we were using were the cause of the issue, we performed another PCR to test the PCR kits we had in stock: [Taq DNA Polymerase](https://www.thermofisher.com/order/catalog/product/EP0401) (Thermo Scientific),
[Phusion](https://www.thermofisher.com/se/en/home/brands/thermo-scientific/molecular-biology/thermo-scientific-pcr/high-fidelity-dna-polymerases-master-mixes-thermo-scientific.html?ef_id=Cj0KCQjwrojHBhDdARIsAJdEJ_fcgpK29q5a43-tUBU6SnjsQmrvP_9zeTUXR1UJxRLUSy6pEt9P4wAaAnikEALw_wcB:G:s&s_kwcid=AL!3652!3!776811043426!e!!g!!phusion%20polymerase!8262701709!187013327995&cid=bid_mol_pch_r01_co_cp1358_pjt0000_bid00000_0se_gaw_bt_pur_con&gad_campaignid=8262701709) (Thermo Scientific),
[DreamTaq DNA Polymerase](https://www.thermofisher.com/order/catalog/product/EP0703) (Thermo Scientific). We performed the PCR using primers and plasmid DNA, coding for the chromoprotein ScOrange, which were kindly provided by Prof. Anthony Forster. The results (Figure 1) showed that all of our kits were working as intended.

![](https://static.igem.wiki/teams/5749/engineering/image1.webp)

**Figure 1** Testing PCR reagents. PCR was done using primers for ScOrange and using plasmid coding for the ScOrange chromoprotein as a template. + ScOrange template added, - no template control.

We then investigated if the issues with the PCR could be due to low quality cell lysates. We theorized that perhaps boiling the cells in EDTA was not enough to lyse cc-1690 strain cells, which have a thick, glycoprotein-rich cell wall, meaning that the DNA was not being released. We decided to try lysing both cc-1690 and cc-3403 (cell wall free strain) using both the previously mentioned protocol, as well as using a different protocol, which we adapted from an RNA prep protocol by Craig D. Amundsen (1999). [[2]](ref2) Here proteinase K is added to the reaction mixture to degrade proteins and inactivate nucleases, as well as SDS, which helps to disrupt cell membranes and denature proteins. However, we feared that SDS could potentially interfere with the DNA polymerase during PCR, so we decided to also try using a SDS-free lysis protocol, which we adapted from a genomic DNA purification with proteinase K protocol by Thermo Fisher Scientific.

For this PCR round we decided to use the same primer pair (PGK-F1/PGK-R1) as used in the cPCR protocol by Nouemssi et al. (2020)[[1]](ref1), which is specific for the nuclear phosphoglycerate kinase (PGK) gene. We hoped that the PGK primers could serve as a positive control for future cloning attempts, as we could not be sure that the primers, which were designed by our team, were working as intended. The reaction was done using Taq DNA Polymerase (Thermo Scientific), following the manufacturers instructions. We once again did not see any product on the gel, meaning that PGK was not being amplified, regardless of which lysis method or strain was used.

We then tried to adapt the PCR for the high CG content of the *C.reinhardtii*. We first tried to follow the user manual for the Taq Polymerase, which suggested that in such cases up to 10% of DMSO could be added to the reaction mixture, as well as extending the denaturation time up to 3 min. We did another PCR, using the same primers (PGK-F/PGK-R) and the same cc-1690/cc-3403 lysates as in the previous try, but with these changes, however it again was unsuccessful (Figure 2).

At the same time, we also tried to perfectly recreate the cPCR protocol from Nouemssi et al. (2020) [[1]](ref1), as we were struggling to understand why we were not seeing PGK being amplified as described in the article, even when using the same primer pair. And for the first time, we were successful in amplifying a *C.reinhardtii* gene sequence. The main difference between this and previous attempts was that the extension temperature was lowered to 68 °C (we previously set the extension temperature at 72 °C, as recommended by the manufacturer). PGK was successfully amplified from both cc-1690 and cc-3403 lysates (figure 2, right side of the gel), although the band for cc-1690 was stronger. We decided to go on using the cc-1690 lysate.

![](https://static.igem.wiki/teams/5749/engineering/image3.webp)

**Figure 2** Optimizing PCR for high GC content. PCR was done using primers for PGK and cc-1690/cc-3403 strain lysates. Different lysis buffers and conditions were used to prepare the lysates: 1 - SDS + proteinase K, 2 - tween-20 + proteinase K, 3 - EDTA only, - no template control. On the right side of the gel, PCR was done using the same PCR program as used in Nouemssi et al. (2020).

Another factor that was considered was the DMSO amount. While DMSO is a great additive to increase PCR yield, it can also interfere with DNA polymerase activity. We decided to reduce the DMSO amount in the reaction mixture to 3%. We tried to run the PCR with the primer pairs for FUS1 (FUS1-F/FUS1-R), MAR1 (MAR1-F/MAR1-R), GP1 (GP1-F/GP1-R) and used PGK (PGK-F/PGK-R) as a positive control. We again were not able to amplify the target genes. However, based on PGK, we can clearly see that the addition of 3% DMSO did appear to greatly improve the PCR yield (Figure 3).

![](https://static.igem.wiki/teams/5749/engineering/image4.webp)

**Figure 3** Cloning attempt #2. We attempted to amplify GP1 (G), FUS1 (F), MAR1 (M) from cc-1690 lysate. PGK was used as a positive control. Taq DNA polymerase was used and 3% DMSO added to the reaction mixture.

Finally, we decided to check if using a different polymerase could improve the results. We decided to try using Phusion (Thermo Scientific) high fidelity DNA polymerase, as the manufacturers claim that it is better suited for high GC templates. We used the GC buffer included in the kit, 3% DMSO was also added into the solution. All of these changes led to a much higher yield (Figure 4). Unfortunately, it also resulted in much lower specificity and with the amount of off-target products, it is impossible to say if we were successful in amplifying our actual targets. This might be caused by the low primer annealing temperature or improper primer design, which we hope to adjust in the future.

![](https://static.igem.wiki/teams/5749/engineering/image2.webp)

**Figure 4** Cloning attempt #3. We attempted to amplify FUS1, MAR1, GP1 from cc-1690 and cc-3340 lysates. Phusion DNA polymerase was used, together with the GC buffer and 3% DMSO added to the reaction mixture. 1 - cc-1690 lysate, 2 - cc-3340 lysate, - no template control.

We conclud, that while cloning *C.reinhardtii* genes is without a doubt **difficult**, some changes can be implemented in order to
greatly improve the results:

1. We did not see a noticeable difference between PCR efficiency when using cell wall free strain versus cell wall intact strain lysates, which shows that our lysis protocol can be used for both. But we did see a major difference between using lysates prepared from fresh, alive cell cultures, as opposed to from old cell samples.
2. Have a good positive control. This will make it much easier to adjust the PCR conditions and to troubleshoot any issues.
3. Try out different polymerases. Standard Taq polymerase can be used for simple applications, but it might struggle with more difficult amplicons. In our case, we found Phusion polymerase to be a good alternative.
4. Addition of DMSO to the reaction mixture can greatly increase the PCR yield for high GC DNA templates. It is best to start out with low amounts (2-3 %) and increase if needed. Keep in mind that this will decrease the primer melting temperature, and the annealing temperature will need to be adjusted accordingly. DMSO also reduces the effectiveness of the DNA polymerase, as well as changing the optimal MgCl2 concentration.

### Primers Used

| Target | Primer | Direction | Sequence (5′→3′) |
| --- | --- | --- | --- |
| **Fus1** | | | |
| Fus1 | Fus1-F1 | Forward | 5′→3′`CCTACGGCCACAACATCACAAC` |
| Fus1 | Fus1-R1 | Reverse | 5′→3′`TGGCTTGTGTCTGAGCATAC` |
| **Mar1** | | | |
| Mar1 | Mar1-F1 | Forward | 5′→3′`GTGAAGATGGCGCGGATGCT` |
| Mar1 | Mar1-R1 | Reverse | 5′→3′`GCGGTGGGGAAGCCACACACC` |
| **PGK** | | | |
| PGK | PGK-F1 | Forward | 5′→3′`AGTCCGAACAACCCACTTAC` |
| PGK | PGK-R1 | Reverse | 5′→3′`CAGAGCTCAGGAGGTGAAATAG` |
| **GP1** | | | |
| GP1 | GP1-F1 | Forward | 5′→3′`TGAGCTCGAGCAGTGCGTACCTGGCGGTATC` |
| GP1 | GP1-R1 | Reverse | 5′→3′`CAGTGTCGACTGGGCTTGGGCATCCAGC` |
| GP1 | GP1-F2 | Forward | 5′→3′`ACTGACGTTAATTAATCAGTGCGTACCTGGCGGTATCTTC` |
| GP1 | GP1-R2 | Reverse | 5′→3′`AACATCGATCGAGGTCTTGGGCTTCTTGTTG` |
| GP1 | GP1-R3 | Reverse | 5′→3′`GAGCCACCCAGATCTCCGTTAACCTTCAGGTTGAGGCCGACCG` |

### References

1. [Nouemssi et al. (2020) - Rapid and Efficient
   Colony-PCR for High Throughput Screening of Genetically Transformed *Chlamydomonas reinhardtii*](https://doi.org/10.3390/life10090186)
2. [Amundsen (1999)
   - Chlamy RNA Prep](https://www.chlamycollection.org/methods/craigs-chlamy-corner/rna-prep/)

{% endblock %}

{% extends "layout.html" %}
{% block title %}Experiments{% endblock %}
{% block lead %}{% endblock %}
{% block page\_content %}

[*E. coli* protocols](#ecoli)
[*C. reinhardtii* protocols](#chlamy)
[References](#refs)

## *Escherichia coli* Protocols

## DNA Protocols

## DNA Resuspension

Plasmids and DNA fragments ordered from synthesis companies often come in powder form in small tubes.
They need to be resuspended before experimental use

### Materials

* Dry DNA
* Tris-HCl ph 8.0 buffer

### Procedure

1. First, given the weight of DNA received, n, use n = cv
   to calculate the required volume of liquid, v, to add to in order to achieve the desired
   final working concentration, c
   * This is usually 10ng/μL, but vary based on the use case.
2. Briefly centrifuge the tube to ensure that all of the dry DNA is at the bottom.
3. Add the required volume v of Tris-HCl ph 8.0 buffer to the tube.
   * It is possible to resuspend DNA in ddH2O but this is not recommended for long term storage.
4. Gently mix by pipetting up and down a few times to ensure all the DNA has been dissolved.
5. Store the resuspended DNA solution at -20°C.

## Competent Cells

Competence is the ability for a cell to take up exogenous DNA. *E. coli* have a low degree of natural competence
but can be treated with CaCl2 to make their cell walls more permeable and increase their competence.

### Materials

* Liquid ON culture
* SOB medium
* Ice-cold 0.1M CaCl2
* Ice-cold 0.1M CaCl2/20% glycerol

### Procedure

1. Take a 500μL aliquot from an overnight culture and add it into 49.5 mL of SOB medium (1:100 dilution).
2. Grow the culture at 37°C with shaking until it reaches an OD600 of 0.4.
3. Place the culture on ice for ~15 minutes, swirling occasionally.
4. Pour the culture into a 50mL Falcon™ tube.
5. Centrifuge at 3500 rpm for 5 minutes at 4°C.
6. Carefully remove as much supernatant as possible without disturbing the pellet.
7. Resuspend the pellet in 100μL ice-cold 0.1M CaCl2 using a sterilized loop.
8. Add 15mL of ice-cold 0.1M CaCl2 and mix gently by pipetting up and down.
9. Incubate the cells on ice for 30 minutes.
10. Centrifuge again at 3500 rpm for 5 minutes at 4°C.
11. Resuspend the pellet in 2mL of ice-cold 0.1M CaCl2/20% glycerol solution.
12. Incubate for 45 minutes on ice.
13. Pipette 50μL aliquots into chilled 1.5mL tubes.
    * Mix gently during aliqouting as cells can settle.
14. Snap freeze any aliquots not required for immediate use in liquid nitrogen and store at -80°C.

* Cells cannot be refrozen once thawed, as this kills them.

## Digestion

### Digestion

#### Materials

* DNA: Plasmid (vector) or gene of interest, variable ng (see formula below for desired molar amount).

copies = (ng × (6.022 × 1023)) / (length × 109 × 650)

* Restriction buffer (10×): 5 µL (50 µL reactions) or 2.5 µL (25 µL reactions).
* Compatible restriction buffer: e.g., Smart Buffer (NEB) or FastDigest Buffer (Thermo Fisher Scientific).
* Restriction enzymes: 1 µL each (double digest) or 2 µL (single digest).
  + Use the appropriate restriction enzymes for your cloning strategy.
* ddH2O: adjusted to the final volume.
* Total reaction volume: 50 µL (or 25 µL for half-volume reactions).

**General note:** Perform all steps on ice unless otherwise stated.

#### DNA Preparation

1. Add the required amount of DNA to reach the desired ng (see formula above).
2. Adjust volume with ddH2O, accounting for the addition of 1 µL of each enzyme:
   * 50 µL reactions → 43 µL ddH2O.
   * 25 µL reactions → 20.5 µL ddH2O.
3. Add 10× reaction buffer (any compatible restriction buffer may be used):
   * 50 µL reactions → 5 µL buffer.
   * 25 µL reactions → 2.5 µL buffer.
4. Do not reduce enzyme volume for smaller reactions.
5. For a double digest: Add 1 µL of each restriction enzyme for both 50 µL and 25 µL reactions (2 µL total).
6. For a single digest: Add 2 µL of restriction enzyme to both 50 µL and 25 µL reactions.
7. Gently tap or flick tubes to mix.
8. Incubate at 37 °C for 30 min.
9. Heat-inactivate enzymes at 80 °C for 20 min.
10. Store the samples at −20 °C.

#### Sample Loading for Gel Electrophoresis

In order to be visible on the gel, load a minimum of approximately 125 ng of DNA in each well on a 1% agarose gel.

#### Fragment Recovery

* Excise and save the plasmid (vector) and gene of interest fragment for gel purification (see Gel Extraction Purification protocol).
* Freeze samples at −20 °C.

## Ligation

### Ligation

#### Materials

* DNA of interest.
* T4 DNA ligase.
* 10× T4 DNA ligase buffer.
* ddH2O.

#### Procedure

1. Label all tubes clearly to distinguish digested samples and controls.
2. Determine plasmid-to-insert ratios based on molarity. For the desired number of copies of plasmid and gene of interest (GOI) added to each reaction, see the formula in the Digestion protocol based on DNA amounts in ng and the number of base pairs. The desired DNA volume for each reaction can be determined using the formula:
   *volume = n(mol) / c(mol/dm3)*
3. The plasmid should be pre-digested with appropriate restriction enzymes leaving restriction cuts compatible with the GOI sequence.
4. Prepare corresponding control tubes: one without DNA and one without ligase. Substitute the missing component with ddH2O.
5. For a 20 µL reaction, add 2 µL of 10× ligase reaction buffer (T4 DNA ligase buffer) to each appropriate tube.
6. Add 1 µL of T4 DNA ligase to each tube.
7. Adjust with ddH2O to reach a final reaction volume of 20 µL.
8. Gently mix by tapping or flicking.
9. Incubate at 22 °C for 30 minutes.
10. Heat-inactivate ligase at 80 °C for 20 minutes.
11. Store samples at −20 °C until further analysis.
12. Run samples on a 1% agarose gel for visualization and comparison of ligation and control reactions (if needed), and see the Transformation protocol.

## Transformation

Transformation is the process of introducing exogenous DNA to a cell. Transformed *E. coli* are
an effective way to replicate and store DNA

### Materials

* Competent cells
* Plasmid of choice
* SOB medium

### Procedure

1. Incubate 50μL competent cells on ice for 15 minutes
2. Add the ~5μL of plasmid to competent cells
   * This may vary slightly depending on the concentration of plasmid DNA
3. Incubate on ice for a further 30 minutes
4. Heat shock the cells at 42°C for 45 seconds
5. Incubate on ice again for 5 minutes
6. Add 950μL of SOB media to the cells
7. Incubate at 37°C for 90 minutes
8. Centrifuge cells for 5 minutes at 4000rpm to pellet
9. Remove 900μL of supernatant, taking care not to disturb the pellet
10. Resuspend the pellet in the remaining 100μL and plate

## Re-streaking

### Re-streaking

1. Take one colony from a plate using an inoculation loop and smear it out on one edge of a new plate.
2. Draw one line vertically through the first lines to the centre of the plate.
3. Sterilize the loop and then go back and forth through the vertical line all the way to the other edge of the plate.

## Overnight Cultures

### Overnight Cultures

#### Materials

* LB media
* Antibiotic of choice (stock solution)
* Plate with grown colonies

#### Procedure

1. Prepare growth media by mixing 5 ml of LB media with adequate amount of antibiotics to obtain a 1× antibiotic concentration.
2. Take one colony from a plate and resuspend it in the growth medium.
3. Incubate at 37 °C, 110 rpm for at least 8 hours (overnight).

## Agarose Gel Electrophoresis

Gel electrophoresis is a technique used to separate DNA fragments by size.
An electric field is applied accross the gel, which causes the negatively charged DNA molecules to move
toward the positive electrode. The agarose matrix slows the movement of the DNA through the gel; smaller molecules
move faster, while larger molecules move slower.

### Materials

* Agarose powder
* 1x Tris/Borate/EDTA (TBE) buffer
* 1x SYBR™ Safe DNA Gel Stain
* DNA sample of choice
* 6x DNA gel loading dye

**Preparation of 1% Agarose Gel (150mL)**

1. Add 1.5g of agarose powder to an Erlenmeyer flask
2. Add 150ml of 1x Tris/Borate/EDTA (TBE) buffer to the flask and swirl to mix
3. Microwave until the agarose dissolves
   * Take care when handling the flask as it will be hot; agar dissolves at 85-90°C
4. Let the flask cool until it can be safely touched
5. Add 1x SYBR™ Safe DNA Gel Stain to the solution and swirl to ensure it is fully mixed
6. Insert the comb into the casting tray
7. Pour the gel solution into the casting tray
8. Gently tap the tray on the bench a few times or use a clean pipette tip to carefully pop any air bubbles
9. Once the gel has solidified, it is ready for use

**Electrophoresis**

1. Ensure that the gel is oriented correctly in the chamber, with the sample wells nearest
   to the cathode (negative electrode)
2. Pour in 1x TBE buffer until the gel is fully covered
3. Prepare the sample mixes in separate Eppendorf tubes or on a clean piece of Parafilm
   * Mix 5μL of sample DNA with 1μL of 6x DNA gel loading dye
4. Pipette the 6μL sample mixes into separate wells on the gel
5. Place the lid and power supply leads onto the electrophosis unit
6. Turn on the power supply and run the gel
   * The gels can be run anywhere in the range of 20-150 volts
   * Higher voltages give shorter run times but may carry the risk of melting the gel

## Gel Extraction Purification

Purify DNA fragments from agarose gels using the Thermo Scientific GeneJET™ Gel Extraction Kit[[4]](#ref4). Optional: Add 100% isopropanol to improve recovery for small or large fragments.

#### Materials

* Agarose gel slice containing DNA fragment of interest (≤ ~1 g)
* Binding Buffer (provided)
* Wash Buffer (provided; supplemented with ethanol)
* Elution Buffer (provided; 10 mM Tris-HCl, pH 8.5)
* GeneJET purification column + collection tube
* Microcentrifuge tubes (1.5 mL)
* Pipettes and tips
* 96–100% ethanol (molecular biology grade)
* (Optional) 3 M sodium acetate, pH 5.2
* Microcentrifuge capable of >12,000 × g

#### Procedure

1. Excise the gel slice containing DNA using a clean scalpel or razor blade.
2. Place gel slice in a pre-weighed tube and add an equal volume of Binding Buffer (1:1 weight-to-volume).
3. Incubate at 50–60°C for 10 min until gel is completely dissolved, mixing occasionally.
4. (Optional) For DNA ≤500 bp or >10 kb, add equal volume of 100% isopropanol and mix.
5. Transfer up to 800 μL of solution to the purification column and centrifuge at >12,000 × g for 1 min. Discard flow-through.
6. Wash column with 700 μL of Wash Buffer and centrifuge at >12,000 × g for 1 min. Discard flow-through.
7. Centrifuge column empty at >12,000 × g for 1 min to remove residual buffer.
8. Transfer column to a clean tube. Add 50 μL Elution Buffer to center of membrane and centrifuge at >12,000 × g for 1 min.
9. Discard column and store purified DNA at −20°C or use immediately in downstream applications.

## Media Protocols

## LB Media

Luria-Bertani (LB) Media, also known as Lysogeny Broth is a nutritionaly
rich solution that can be used to grow bacterial cultures. LB Media typically contains peptides, carbohydrates, vitamins,
and minerals which are needed by bacteria. LB can be prepared by mixing its constituent
components, or from commerically available powder mixes.

#### Materials

* NaCl
* Bacto tryptone
* Yeast extract
* NaOH
* ddH2O

#### Procedure

1. In a 1L bottle, add NaCl (0.17M), Bacto tryptone (1% w/v), yeast extract (0.5% w/v),
   ddH2O to 600mL, 100µL of 5M NaOH.
2. Autoclave for 20 min within 2h, store in the fridge.

## LB Agar

LB agar is LB media with added agar. The agar sets the LB into a gel,
which provides a solid media for bacterial colonies to grow on. Antibiotics can be
added to the gel before pouring onto plates, which can then be used to screen bacteria for
antibiotic resistance. LB Agar can be made by adding agar powder while preparing LB media or
by from commercially available powder mixes.

#### Materials

* LB media
* Agar powder
* Antibiotic of choice (1000x)
* 30 petri dishes

#### Procedure

1. In a 1L bottle, add 600mL of LB media and 9g of agar and shake.
2. Autoclave for 20 min within 2h.
3. Let it cool until you can touch the bottle without burning yourself and add 600 µL of
   1000x antibiotics of choice.
4. Gently mix, pour into petri dishes (20mL per dish) and let it solidify for an hour at
   room temperature. Once solid, turn upside down the dishes at room temperature for a few
   hours, then store in the fridge.

## SOB Media

Super Optimal Broth (SOB) media is a nutritionaly rich solution that is optomised for
the preparation and transformation of competent cells. SOB is an adjusted version of LB media.
SOB can be prepared by mixing its constituent components, or from commerically available powder mixes.

#### Materials

* Yeast extract
* Bacto tryptone
* NaCl
* KCl
* NaOH
* ddH2O

#### Procedure

1. In a 1L bottle, add yeast extract (0.5% w/v), Bacto tryptone (2% w/v), NaCl (10mM), KCl
   (2.5mM), 600mL of ddH2O, 120µL 5M NaOH.
2. Autoclave for 20 min within 2h, store in the fridge.

## Cultivation

## Overnight cultivation

#### Materials

* LB media
* Antibiotic of choice (stock solution)
* Plate with grown colonies

#### Procedure

1. Prepare growth media by mixing 5ml of LB media with adequate amount of antibiotics to
   obtain a 1x antibiotic concentration.
2. Take one colony from a plate and resuspend it in growth medium.
3. Incubate at 37°C, 110rpm for at least 8 hours (overnight).

## Re-streaking

1. Take one colony from a plate using an inoculation loop and smear it out on one edge of a
   new plate.
2. Draw one line vertically through the first lines to the centre of the plate.
3. Sterilize the loop and then go back and forth through the vertical line all the way to
   the other edge of the plate.

## Digestion

#### Materials

* DNA: Plasmid (vector) or gene of interest, variable ng (see formula below for desired
  molar amount).

copies = (ng \* (6.022 \* 1023)) / (length \* 109 \* 650)

* Restriction buffer (10x): 5 µL (50 µL reactions) or 2.5 µL (25 µL reactions).
* Compatible restriction buffer: e.g., Smart Buffer (NEB) or FastDigest Buffer
  (Thermo Fisher Scientific).
* Restriction enzymes: 1 µL each (double digest) or 2 µL (single digest).
  + Use the appropriate restriction enzymes for your cloning strategy.
* ddH2O: adjusted to the final volume.
* Total reaction volume: 50 µL (or 25 µL for half-volume reactions).

General note: Perform all steps on ice unless otherwise stated.

#### DNA Preparation

1. Add the required amount of DNA to reach the desired ng. (see formula above).
2. Adjust volume with ddH2O, accounts for addition of 1 µL of each enzyme:
   * 50 µL reactions → 43 µL ddH2O.
   * 25 µL reactions → 20.5 µL ddH2O.
3. Add 10x reaction buffer, any compatible restriction buffer may be used:
   * 50 µL reactions → 5 µL buffer.
   * 25 µL reactions → 2.5 µL buffer.
4. Do not reduce enzyme volume for smaller reactions.
5. For a double digest: Add 1 µL of each restriction enzyme for both 50 µL and 25 µL
   reactions (2 µL total).
6. For a single digest: Add 2 µL of restriction enzyme to both 50 µL and 25 µL reactions.
7. Gently tap or flick tubes to mix.
8. Incubate at 37 °C for 30 min.
9. Heat-inactivate enzymes at 80 °C for 20 min.
10. Store the samples at −20 °C.

#### Sample loading for gel electrophoresis

In order to be visible on the gel, load a minimum of ~125 ng of DNA in each well on a 1% agarose gel.

#### Fragment recovery

* Excise and save the plasmid (vector) and gene of interest fragment for gel purification
  (See Gel Extraction Purification protocol).
* Freeze samples at −20 °C.

## Ligation

Ligation

#### Materials

* DNA of interest.
* T4 DNA ligase
* 10x T4 DNA ligase buffer
* ddH2O

#### Procedure

1. Label all tubes clearly to distinguish digested samples and controls.
2. Determine plasmid-to-insert ratios based on molarity. For the desired amount of copies
   of plasmid and gene of interest, GOI, added to each reaction, see the formula in
   Digestion protocol based on DNA amounts in ng and the amount of basepairs of the DNA.
   The desired amount of volume of DNA added to each reaction can be determined using the
   volume formula below.
   volume = n(mol) / c (mol/dm3)
3. The plasmid should be pre-digested with appropriate restriction enzymes leaving
   restriction cuts compatible with the GOI sequence.
4. Prepare corresponding control tubes, one without DNA, and one without ligase. Both
   substituted with ddH2O.
5. For 20 µL reaction, add 2 µL of 10x ligase reaction buffer (T4 DNA ligase buffer) to
   appropriate tubes.
6. Add 1 µL of T4 DNA ligase to each tube.
7. Adjust with ddH2O to reach a final reaction volume of 20 µL.
8. Gently mix by tapping or flicking.
9. Incubate at 22 °C for 30 min.
10. Heat-inactivate ligase at 80 °C for 20 min.
11. Store samples at −20 °C until further analysis.
12. Run samples on a 1% agarose gel for visualization and comparison of ligation and
    control reactions if needed and see Transformation protocol.

## Gibson Assembly

#### Materials

* GeneJET™ PCR Purification Kit
* Assembly Master Mix
* ddH2O

#### Procedure

1. PCR the individual gene fragments to include overlapping region.
2. Purify the gene fragments with PCR purification kit according to the GeneJET™ PCR
   Purification Kit User Guide [[5]](#ref5).
3. Mix the gene fragment(10–100 ng of each ~6 kbp DNA fragment) with 10 μL of Assembly
   Master Mix. Add ddH2O to reach final volume of 20μL.
4. Incubate at 50°C for 12 min.
5. Transform to competent cells following the transformation protocol.

## Protein Protocols

## Lysis

To extract the produced protein from the cell, we rupture the cells with lysozyme.

### Materials

* 1 M Tris-HCl, pH 8.0
* 0.5 M EDTA, pH 8.0
* NaCl
* Triton X-100
* Lysozyme stock solution (10 mg/mL in 10 mM Tris-HCl, pH 8.0)
* BL21 Z17-induced samples

### Procedure

1. Prepare the lysis buffer with the following composition:
   * 10 mM Tris-HCl, pH 8.0
   * 1 mM EDTA
   * 100 mM NaCl
   * 0.5% (v/v) Triton X-100
2. Resuspend the cell pellet in **300 µL** of lysis buffer.
3. Add **25 µL** of lysozyme stock solution.
4. Mix thoroughly by vortexing for a few seconds.
5. Incubate the samples at **37 °C for 30 minutes**.
6. Centrifuge the samples at **maximum speed** for **3 minutes**.
7. Carefully collect the supernatant, which contains the soluble protein fraction.
8. To obtain data about the protein content of the cell perform SDS-page gel

## Urea Extraction

Since the proteins expressed were insoluble, and were not present in a supernatant after cell
lysis, we have performed Urea denaturation to solubilize and purify the protein

### Materials

* Urea powder (for preparing 4 M, 6 M, and 8 M solutions)
* Deionized or distilled water
* Sample pellets (from 2 mL bacterial culture)

### Procedure

1. Prepare urea solutions of **4 M**, **6 M**, and **8 M** concentrations.
2. Add **200 µL** of the appropriate urea solution to each sample.
   Note: Since the samples contain only pellets from 2 mL of culture, 200 µL was used instead of the original Costa Rica protocol volume.
3. Incubate the samples for **60 minutes at room temperature**.
   Observation: The first incubation of 35 minutes was insufficient; 60 minutes gave better results.
4. Centrifuge the samples at **5000 × g for 5 minutes**.
5. Carefully collect and store the supernatant. Proceed to the next urea concentration.
6. Repeat steps 2–5 until the samples have been treated with the highest (8 M) urea concentration.
   Note: When urea concentraiton at which Pretein denatures is known,
   just perform one step at given urea concentration
7. Run an **SDS-PAGE gel** using **10 µL** of each sample.
8. Electrophorese the gel at **100 V for 45 minutes**.

## Refolding

The protocol has been recomended by Cytiva [[2]](ref2)

### Materials

* Urea powder (for preparing 8 M, 6 M, 5 M, 4 M, 3 M, 2 M, 1 M, and 0 M buffers)
* Imidazole (prepare 5 mM solution; 0.023828 g per 20 mL)
* Binding buffer (base solution, modified by adding urea and imidazole)
* Elution buffer (Cytiva HisTrap HP protocol base, with added urea)
* ddH₂O (filtered, non-pyrogenic “Steril-R”)
* 20% ethanol (for column storage)
* HisTrap HP column (1 mL)
* Protein sample (urea-extracted)
* Microcentrifuge tubes (1.5 mL and 2 mL)
* 15 mL Falcon tubes
* Pipettes, tips, and syringes (for buffer loading and washing)

### Procedure

1. Prepare the buffers according to the Cytiva protocol [[2]](ref2)

and prepare the prepare
5ml each of running buffer at a decreasing urea concentration- Wash the HiTrap HP column with 15 mL ddH₂O (filtered, non-pyrogenic “Steril-R”) to remove ethanol.
- Wash with 5 mL binding buffer.
- Perform a blank run:
  * 5 mL ddH₂O
  * 5 mL elution buffer
  * 10 mL binding buffer
- Prepare the protein sample by mixing 0.5 mL of protein solution with 1 mL binding buffer and load it into the column.
- Wash with 10 mL binding buffer.
   NOTE: for refolding run the colum with 5ml
  of each runnign buffer at decreasing concentraiton of urea
- Elute and collect with 5 mL elution buffer.
- Clean the column with 5 mL ddH₂O followed by 5 mL binding buffer.
- Run 5 mL of 20% ethanol through the column for storage.
- Replace upper and lower stoppers.

## Expression

To express a prtein BL21 strain of  *E. coli* has been used. in this strain protein produciton is induced by IPTG.

## Materials

* LB medium (Lysogeny Broth)
* Kanamycin (antibiotic stock solution, typically 50 mg/mL)
* IPTG (Isopropyl β-D-1-thiogalactopyranoside, 1 M stock)
* *E. coli BL21* strain (containing required plasmid)

## Procedure

The steps of this protocol are devided into 3 sections for better oritentation

### Initiation

1. Prepare **50 mL LB medium** in a Erlenmeyer flask.
   Add Kanamycin (50 µL of stock) and mix thoroughly.
2. Add **500 µL** of an overnight culture (we used *E. coli* BL21).
   Alternatively use a single colony from a plate (overnight culture is preferred for faster induction timing).
3. Incubate the flask at **37 °C** with shaking at
   **100 RPM**.
4. Monitor growth by measuring OD600. Grow until OD600 is between **0.6 – 0.9**. Take measurements hourly and increase frequency to every 30 minutes as you approach the target range.
5. When the culture reaches the target OD, divide the remaining culture into two flasks: one will be induced and the other left as a control.
6. Induce the experimental flask by adding IPTG to a final concentration of **1 mM**. (Leave the control flask uninduced.)

### Measurement

1. After addition of IPTG, the samples (IPTG induced and control/non-induced) were left for
   a total of 5h at 37 °C at 100 RPM (samples can be taken even at 3h after induction).

### Sample collection

1. Spin the Eppendorf tubes in 6000rpm in 5 min and pour of the supernatant.
   If you cant get all of the supernatant out, spin it down again and carefully pipet it out
   . freeze at -20 °C.
2. For the remaining culture in the e-flasks, add to Eppendorf tubes (aliquot 2ml),
   centrifuge down (6000RPM at 5 min) and pour off supernatant (pipet if required),
   freeze at -20 °C.

## SDS Page

o Evaluate the protein content of cells we prepare samples and load on denaturing SDS-page gel

### Materials

* Frozen cell samples
* ddH2O (deionized/distilled water)
* Tris-Glycine SDS sample buffer
* β-mercaptoethanol (BME)
* Protein ladder (molecular weight marker)
* SDS-PAGE gel (precast or hand-cast)
* Electrophoresis buffer (Tris-Glycine-SDS running buffer)

### Procedure

1. Resuspend the samples in **50 µL** of ddH2O.
2. From each sample, take **10 µL** and add:
   * **9.5 µL** of Tris-Glycine SDS sample buffer
   * **0.5 µL** of β-mercaptoethanol (BME)
3. Mix gently and boil the samples for **5 minutes at 95 °C**.
4. Assemble the SDS-PAGE gel into the electrophoresis tank and fill with SDS buffer.
5. Load your prepared samples into the wells.
6. Load **5 µL** of protein ladder
   (we used [Broad Range Protein Molecular Weight Markers](https://se.promega.com/products/cloning-and-dna-markers/dna-ladder-rna-ladder/broad-range-protein-molecular-weight-markers/?catNum=V8491) from promega) into one well as a molecular weight reference.
7. Run the gel at **100 V** (the current reached ~50 mA) until the dye front has migrated slightly beyond **¾ of the gel length**.

## *Chlamydomonas reinhardtii* Protocols

## DNA Protocols

## Electroporation

Electroporation uses short, high voltage pulses to create temporary pores in the cell
membrane, allowing exogenous DNA to enter the nucleus.

**Day 1**

1. Spin down 1mL of algae solution in a Eppendorf tube and remove the supernatant
   * This should be done at a low RPM to avoid killing the cells!
2. Resuspend cells in algal wash buffer to get a final concentration of 1×108 cells/mL
3. Add 15ng/kb of linear DNA to the cuvette
   * Linear DNA should be sufficiently concentrated so at most 50µL are added
4. Add 1×107 cells to a 2mm electroporation cuvette (100μL of the concentrated cells)
5. Insert the cuvette into the electroporator and apply the follwing pulses:
   * These settings are for a 2mm cuvette. If using a different size cuvette, adjust volatages accordingly
   * Voltages should be adjusted in order to generate an equal field strength (in V/cm). This means that voltages should be halved for a 1mm cuvette
6. Two poring pulses of 8ms each, the first at 250V, the second 150V.
   * These pulses porate the cell wall and cell membrane; if using a cell wall deficient strain a
     single poring pulse of 210V is sufficient
7. Five transfer pulses of 50ms each, starting at 20V with a 40% decay rate (pulses of 20, 12, 7.2, 4.3, and 2.6V)
8. Remove the cuvette from the electroporator and let it rest for ca. 15 minutes
9. Add 4mL of TAP media with sucrose 40mM to a 15mL centrifuge tube
10. Carefully transfer the contents of the cuvette to the centrifuge tube
11. Rinse the cuvette with 1mL of the TAP/sucrose mix then transfer to the centrifuge tube for a final volume of 5mL
12. Place the centrifuge tube horizontaly on a shaker and secure with tape
13. Run the shaker overnight at 120rpm on the lowest light setting

**Day 2**

1. Collect cells by centrifuging at 1000rcm for 5 minutes
2. Spread cells ontp TAP agar plates with the appropriate selection antibiotic
   * For our experiments we used Paromomycin 20 μg/mL
3. Leave the plates to grow at room temperature with no shaking, approximately 15-20mE light intensity
4. Transformants typically appear after 5-7 days

**Day 7-9**

1. Once transformants appear, transfer putative antibiotic-resistant colonies to individual liquid cultures

## Gibson Assembly

### Gibson Assembly

#### Materials

* GeneJET™ PCR Purification Kit
* Assembly Master Mix
* ddH2O

#### Procedure

1. PCR the individual gene fragments to include overlapping regions.
2. Purify the gene fragments with the PCR purification kit according to the GeneJET™ PCR
   Purification Kit User Guide [[5]](#ref5).
3. Mix the gene fragment (10–100 ng of each ~6 kbp DNA fragment) with 10 μL of Assembly
   Master Mix. Add ddH2O to reach a final volume of 20 μL.
4. Incubate at 50 °C for 12 minutes.
5. Transform into competent cells following the Transformation protocol.

## Genomic Gene Cloning

To amplify highly repetitive and GC rich C. Reinhardtii genes, we used a modified colony PCR
protocol by Nouemssi et al. (2020)[[3]](ref3) .

### Materials

* *Chlamydomonas reinhardtii* cells (strain: cc-1690)
* 10 mM EDTA solution
* Phusion DNA Polymerase (Thermo Scientific)
* GC buffer (provided with Phusion polymerase)
* DMSO (to a final concentration of 3%)
* PCR reagents and primers specific to the target sequence

### Procedure

1. Using a sterile pipette tip, transfer a small amount of *C. reinhardtii* cells into a microcentrifuge tube.
2. Resuspend the cells in **50 µL of 10 mM EDTA**.
3. Vortex briefly to mix and **boil at 95 °C for 10 minutes** to lyse the cells.
4. Immediately place the samples on **ice for 1 minute**, then vortex again.
5. Centrifuge the samples at **3500 × g for 5 minutes**.
6. Carefully collect **2 µL of the supernatant** — this will serve as the DNA template for PCR.
7. Prepare the PCR reaction mixture using **Phusion DNA Polymerase** with **GC buffer** and **3% DMSO**.
8. Run the PCR with the following thermal cycling conditions:
   * **Initial denaturation:** 98 °C for 30 seconds
   * **34 cycles of:**
     + Denaturation: 98 °C for 10 seconds
     + Annealing: 55 °C for 30 seconds
     + Extension: 72 °C for 2 minutes
   * **Final elongation:** 72 °C for 10 minutes
9. Visualize PCR products on a **1% agarose gel**.

## Media Preparation

## TAP Media

TAP (Tris-Acetate-Phosphate) is the standard growth media for *C. reinhardtii*.
It contains specific salts and trace elements, which allow the cells to grow
both phototrophically (using light as a source of energy) and
heterotrophically (without using light). Our TAP media protocol requires phosphate buffer and
TAP salts, which were provided to us but the are desrcibed below for completeness.
Solid media for plates can also be created by adding agar while preparing TAP media

**Phosphate Buffer (100mL)**

1. In a suitable flask mix:
   * 10.8g Dipotassium phosphate (K2HPO4)
   * 5.6g Monopotassium phosphate (KH2PO4)
   * Distilled water (dH2O) to 100mL

**TAP Salt Buffer (500mL)**

1. In a suitable flask mix:
   * 20g Ammonium chloride (NH4Cl)
   * 5g Epsom Salt (MgSO4·7H2O)
   * 2.5g Calcium chloride, dihydrate (CaCl2·2H2O)
   * Distilled water (dH2O) to 500mL

**TAP Media (1L)**

1. Add 800mL of dH2O to a large graduated vessel
2. Under continuous stirring at the following:
   * 2.42g Tris base (Trizma)
   * 1mL Phoshate buffer
   * 10mL TAP salt buffer
   * 1mL Hutner's trace elements
3. Fill the vessel to 1L with dH2O
4. Check the pH of the solution and adjust to pH7 by adding glacial aecitic acid (usually ~1mL)
5. Transfer to laboratory bottles and autoclave to sterilise
   * If making TAP agar, add 1.5% (w/v) plant agar before autoclaving
   * To make selective media, antibiotic such as paromomycin can be added after autoclaving,
     once the media has sufficiently cooled.

## Culture and Measurement

## Liquid Culture

Liquid cultures can be grown in Erlenmeyer flasks on a shaker. In order to allow efficient
stirring and aeration, the total media volume should not exceed one fifth of the flask volume. Our cultures were
grown under the following conditions:

* **Shaker speed:** 140rpm
* **Light intensity:** 50µmol photons m-2s-1
* **Temperature:** ~20°C
* Cultures were refreshed every 2 weeks by discarding half of the flask contents and replacing
  the discard with the same volume of fresh TAP media.

## Operating Cell Counter

To calculate how many *C. reinhardtii* cells were in a liquid culture we
used a cell counter, the Countess™ II FL Automated Cell Counter from Thermofisher Scientific.
Our experience showed that the cell counter worked best for cell concentrations in the range 1×105-
1×107cells/mL. Because the cell counter calculates a final concentration by extrapolating from the
cells in the viewport, measurements at lower concentrations are extremely sensitive to even small amounts of dust or debris
that may be present on the slide. At higher concentrations, cells can become too densly packed and edge detection becomes
difficult, leading to underestimation. For this reason, it is important to inspect the image before saving the results.

1. Take a sample from a liquid culture of *C. reinhardtii*
2. If the culture is too dense dilute it with TAP media, making note of the dilution factor
3. Immobilize the sample cells with 96% acetic acid, to a final volume of 1/10 of the sample
   * e.g a final sample volume of 50µL should consist of 45µL from the liquid culture and 5µL of acetic acid
4. Prepare a Countess™ reusable haemocytometer slide and cover glass by washing with dH2O and gently drying
   with low lint tissue paper
   * Care should be taken when drying, as the cell counting chambers of the haemocytometer
     have sharp edges which may tear the tissue, leaving debris
5. Load 10µL liquid into the haemocytometer slide, and insert into the cell counter
6. The cell counter captures an image of the viewport, calculates the number of visible cells, and the sample concentration
   * The sample concentration calculation assumes a sample dilution of 1:1 with trypan blue. To account for this,
     the displayed concentration needs to be multiplied by 10/18 (multiply by 10/9 to account for the actual sample dilution
     due to the addition of the aecetic acid, then multiplied by 1/2 to account for the assumed trypan blue dilution)

## Growth Curve

In order to understand the growth dynamics of *C. reinhardtii* it is necessary to plot a growth curve.

1. Using the Cell Counter protocol to measure the initial cell concentration in liquid cultures of interest
2. Dilute the liquid cultures to the same starting value
   * We aimed for an initial starting concentration of 2.5×105cells/mL
3. Use the Cell Counter to measure the concentration every 24 hours
   * To increase accuracy, use multiple technical and biological replicates; use multiple flasks,
     and take measurements form each flask multiple times
4. Plot the measured concentrations against time

## Sedimentation

To determine the efficacy of induced sedimentation, we need to first observe the
natural rate of sedimentation in *C. reinhardtii*, when colonies are left undisturbed.

1. For each colony of interest, mix 5mL of liquid culture with 10mL of TAP media in a Falcon® tube
2. Take an initial sample (t=0) from each tube and measure the concentration
3. Store Falcon® tubes in the refrigerator when not in use, taking care not to agitate the tubes too much when moving them
4. Take further samples every 30 minutes (t=30, 60, 90, 120) and plot the results

## Flocculation

To determine the efficacy of genetically induced flocculation, we need to also observe the
rate of chemically (pH change) induced flocculation in *C. reinhardtii*. This protocol modifies a
protocol by Fan, et al. (2017)[[1]](#ref1).

1. Prepare high pH TAP media by adding 5mM CaCl2 and adjusting to a final pH of 10-11 with NaOH
2. Centrifuge 10mL of liquid *C. reinhardtii* culture at 1000rcm for 2 minutes
3. Remove as much supernatant as possible without disturbing the algal pellet
4. Add 10mL of high pH TAP to the pellet and resuspend
   * Use regular TAP media for the control
5. Measure the concentration initially and at subsequent 10 minute intervals
   * Take measurement samples from the top of the tube (if using a 10mL tube, pipette from the 9mL mark)

## Salt Tolerance

Since the expression of our flocculation genes can be controlled by an salt (NaCl) promoter,
it is important to test how tolerant *C. reinhardtii* is to high salinity cultures.

1. In freshly autoclaved Erlenmeyer flasks, add 2.90 mL liquid TAP media and 10 mL of liquid *C. reinhardtii* culture to each flask.
2. Set up three flasks for each strain of interest
   * In our case: wild type cc-125 and cell wall-deficient strain cc-3403
3. To the flasks with cc-3403, add 100 µL arginine (1:1000 dilution).
   * Arginine is essential for the growth of cc-3403 and is not present in normal TAP media
4. For each strain, add NaCl so that the 3 flasks have the following final concentrations: 0mM (control), 20mM, 100mM
5. Grow the cultures for 5 days under standard conditions.
6. Measure cell counts in duplicate (technical replicates) every 24 hours.

### References

1. [Fan et al. (2017) - Flocculation of *Chlamydomonas reinhardtii* with Different Phenotypic Traits by Metal Cations and High pH.](https://doi.org/10.3389/fpls.2017.01997)
2. [Cytiva (2020) - Rapid and efficient purification and refolding of a (histidine)6-tagged
   recombinant protein produced in E. coli as inclusion bodies](https://cdn.cytivalifesciences.com/api/public/content/digi-16306-pdf)
3. [Nouemssi et al. (2020) - Rapid and Efficient Colony-PCR for High Throughput Screening of
   Genetically Transformed *Chlamydomonas reinhardtii*.](https://doi.org/10.3390/life10090186)
4. [GeneJET™ Gel Extraction Kit User Guide. (PDF)](https://documents.thermofisher.com/TFS-Assets/LSG/manuals/MAN0012661_GeneJET_Gel_Extraction_UG.pdf)
5. [GeneJET™ PCR Purification Kit User Guide. (PDF)](https://documents.thermofisher.com/TFS-Assets/LSG/manuals/MAN0012662_GeneJET_PCR_Purification_UG.pdf)
6. Forster AC, Bao L, Liljeruhm J (2025) Synthetic Biology: a lab manual, 2nd edition. World Scientific, Singapore.

{% endblock %}

{% extends "layout.html" %}
{% block title %}Results from *Escherichia coli* and *Chlamydomonas reinhardtii*{% endblock %}
{% block lead %}In the following section we present all of our results, both successful and unsuccessful, as scientific transparency
is an essential value in research. Although we were unable to complete all of the planned steps,
documenting our full process ensures that future teams can build
upon our progress in developing inducible flocculation in *C. reinhardtii*.{% endblock %}
{% block page\_content %}

[*E. coli* results](#ecoli)
[Purification](#purification)
[Digestion Ligation Transformation](#digestion-ligation)
[*Chlamydomonas* results](#chlamy)
[Growth Curve](#growth)
[Flocculation](#flocculation)
[Electroporation](#electroporation)
[Summary](#summary)
[Outlook](#outlook)
[Appendix](#appendix)

# ***E. coli* results**

## Purification Results

*#### Z17-EhaA overexpression*

After transforming the pET plasmid containing the Z17 and the EhaA autotransporter construct
(hereafter referred to as Z17) into *E. coli* BL21, we proceeded with expression and
isolation of the protein. Figure 1 illustrates successful production of Z17 following IPTG
induction at 37°C. After 2 hours of induction, the Z17 band was undemostrated, but high overexpression
could be observed at both 3 h and 5 h. These results indicate that extending
induction beyond 3 h does not significantly enhance Z17 yield (comparing the strength of the
protein band from background), and future optimization (eg. to enhance solubility, comparing the strength of the protein band from background, and future optimization may therefore focus on induction conditions other than prolonged time, such as induction temperature or IPTG concentration.)

![](https://static.igem.wiki/teams/5749/results/fig-1-bmc-results.webp)

**Figure 1.** SDS-PAGE analysis of *E. coli* BL21 cell pellets containing the Z17 expression plasmid. Protein expression was induced(+) with IPTG and samples were collected at 2h, 3 h and 5 h post-induction, induced and noninduced (-) by IPTG. A prominent band corresponding to Z17 is observed at ~50 kDa.

*#### Optimizing Z17 solubility and purification*

After expressing the Z17 construct (Figure 1), cells were lysed and the supernatant was
subjected to His-tag affinity purification. Ten microliters of the elution fraction were loaded
onto an SDS-PAGE gel. However, no detectable Z17 band was observed in the elution, and since
the flowthrough fractions were not collected, the point of protein loss could not be determined
(data not shown). We hypothesized that the construct either failed to bind to the His-tag
column or was retained in the pellet due to the formation of inclusion bodies.

To address this issue, we first attempted protein expression at a lower temperature to reduce
potential misfolding and aggregation. In parallel, we tested urea denaturation to solubilize
inclusion body–associated protein and enhance His-tag accessibility for purification.

To evaluate the effect of temperature on Z17 expression, we compared protein production at 20 °C
and 37 °C. SDS-PAGE analysis of whole-cell pellets revealed a distinct band at approximately
50 kDa, corresponding to the expected size of Z17. The band was visible at both temperatures
(Figure 2, lane 2; Figure 1, lane 6) but appeared more intense relative to background proteins
at 37 °C, consistent with the optimal growth temperature of *E. coli*.

These findings suggest that higher expression temperatures increase Z17 yield, though possibly
at the cost of solubility. This emphasizes the importance of optimizing induction temperature
to balance protein production and proper folding. Future experiments could include intermediate
temperatures (25–30 °C) to identify the optimal compromise between yield and solubility.

![](https://static.igem.wiki/teams/5749/results/fig-2-bmc-results-new.webp)

**Figure 2.** SDS-PAGE analysis of *E. coli* BL21 cell pellets containing the Z17 expression
plasmid. Protein expression was induced with IPTG and incubated for 3h at either 20°C (lane 2) or
37°C (lane 4). Negative control without IPTG induction is also shown (lane 3). A prominent band at ~50 kDa
corresponds to Z17. Protein Molecular Weight Markers (ThermoFisher) were included for size reference.

*#### Urea denaturation*

After the unsuccessful His-tag purification of lysed Z17, we hypothesized that the His-tag
might be buried within the protein’s folded structure or trapped inside inclusion bodies. To
overcome this, we performed urea denaturation to solubilize the protein and expose the His-tag.
This protocol was adapted from the Costa Rica iGEM 2019 team and optimized for our construct by
treating Z17-containing cell pellets with 8 M urea (lanes 1–5 in Figure 3).

We tested varying incubation times and temperatures to assess their impact on solubilization
efficiency. The results showed that temperature had minimal influence, whereas longer
incubation times increased the solubilization of a wider range of background (non-target)
proteins. Furthermore, the addition of glycine, as suggested by the Costa Rica 2019 team,
did not improve extraction efficiency.

Based on these findings, we selected room temperature and a 1-hour incubation time as the
optimal conditions for urea denaturation (see Urea Extraction protocol). This step allowed us
to proceed with His-tag purification, with the goal of refolding and purifying Z17.
The observed bands indicate the presence of our target protein (~50 kDa) along with other
background proteins, confirming successful denaturation and solubilization (Figure 3).

![](https://static.igem.wiki/teams/5749/results/fig-3-bmc-final.avif)

**Figure 3.** SDS-PAGE analysis of supernatant fractions after urea extraction from *E. coli*
BL21 cells carrying the Z17 plasmid, harvested 5h post-induction. To optimize urea extraction, one
parameter was varied per lane: lane 1, addition of glycine for 1h at 20°C; lane 2, incubation at 37°C for 1h;
lane 3, incubation at 30°C for 1h; lane 4, incubation at 20°C for 2h; lane 5, incubation at 20°C for
3h. The expected Z17 band is at ~50 kDa. Whe have took the overexpressed spind-down cells form Figure 1, lyzed(by lysozyme and freezing)
and solubilized by urea. From this supernatant was loaded to the gel.

*#### Successful His-tag purification*

After expressing Z17 in *E. coli* BL21 and denaturing the protein with 8 M urea, we
performed His-tag purification as an additional purification step. The denatured lysate was
loaded onto the His-tag column, and fractions were collected for analysis. A visible band in
lane 9, Figure 4, corresponding to the first flowthrough, indicated that not all proteins were
retained on the column. This partial loss may be attributed to the slow, manual application of
the sample, which could have reduced binding efficiency compared to using a pump.

Despite this, Z17 was successfully eluted (Figure 4, lane 8). The target protein was primarily
recovered within the first 5 mL of elution, suggesting that future purifications could be
optimized by reducing the total elution volume from 15 mL to 5 mL, thereby increasing protein
concentration and minimizing buffer use.

![](https://static.igem.wiki/teams/5749/results/fig-4-bmc-final.avif)

**Figure 4.** SDS-PAGE analysis of His-tag purification and attempted
refolding of Z17 after urea denaturation and lysis of *E. coli* BL21 cells
induced for 5 h with the Z17-containing plasmid. All samples loaded have passed
through the His-tag column, representing the flow through of the indicated steps.
Protein purification was performed by first loading the sample (lane 9) onto the
column. Elution was carried out using three sequential 5 mL fractions of elution
buffer (lanes 6–8).

## Digestion, Ligation, and Transformation Results

To construct the Z18-TEV-GFP-EhaA plasmid, the pET vector (TEV-GFP-EhaA) and Z18 insert were
digested using BamHI and XbaI. The digestion efficiency was confirmed by agarose gel
electrophoresis, which showed clear band separation at the expected sizes. Prior to this step,
digestion of the pLM005 plasmid had been successfully performed using the same restriction
enzymes, validating both the enzyme compatibility and the workflow.

Initial ligation attempts using the digested pET plasmid and Z18 insert did not yield colonies
after transformation, suggesting possible technical issues in the ligation reaction. To identify
potential causes, a control ligation with pLM005 was performed using identical conditions, which
also failed to produce colonies. This indicated that the issue was not specific to the Z18
construct but likely related to reaction efficiency, possibly due to low plasmid concentration
or reduced ligase activity.

To troubleshoot, reaction parameters were systematically optimized. The insert-to-vector molar
ratio was varied (2:1, 3:1, and 5:1), and the ligation incubation time was extended to 11 hours
at 22 °C. Freshly prepared ligase was also used to minimize the risk of enzyme degradation.
After these optimizations, the ligation reactions were transformed into DH5α and JM109
competent cells.

Successful transformation was confirmed by the presence of colonies on selective media.
The highest colony yield was obtained for JM109 cells transformed with the 5:1 insert-to-vector
ratio, while lower ratios and unspun controls produced significantly fewer colonies.
These results demonstrate that both the increased insert ratio and extended ligation time were
critical for achieving successful plasmid assembly and transformation.

# ***Chlamydomonas* results**

## Growth Curve of *Chlamydomonas reinhardtii*

We began by establishing growth curves for *Chlamydomonas reinhardtii* under our specific lab conditions for the wild type strain
 *CC-125*  and a starch over-accumulating strain dp120 (Figure 5). Cultures were maintained at a light intensity of 50 µmol
photons m-2s-1, shaken at 140 rpm, and kept at approximately 20°C. The cell concentrations were measured using a cell counter. As the cell counter has a large error rate, 4 samples were measured for each of the algae strains. The unexpected peak in cell concentration observed on day
4 suggests that the measurements may not accurately reflect true cell concentration. We suspect that rapid cell sedimentation was the
main cause of these unexpected results. Although the resulting growth curve does not provide a reliable model of *C. reinhardtii*  growth, this experiment was
an important step in learning how to culture this organism, that all of the team members but one were previously unfamiliar to,
and troubleshoot cell count measurements. For future growth curve experiments,
we emphasize the importance of thoroughly homogenizing culture samples prior to measurement.

![](https://static.igem.wiki/teams/5749/results/growth-curve.webp)

**Figure 5.** Seven day growth curve of *C. reinhardtii* strains wild type CC-125 and starch over-accumulating strain
dp120 measured in duplicates. The cells were cultured under a light intensity of 50 µmol photons m-2s-1, a rotation of
140 rpm and a temperature of approximately 20°C.

## Flocculation Efficiency

To test our project concept, we compared natural sedimentation of *C. reinhardtii* cells with chemically induced flocculation.
By adding NaOH and CaCl2 to the growth medium, we observed aggregation of algal cells followed by accelerated sinking.
Flocculated cells settled at the bottom of the container significantly faster than untreated cultures.
These results validated our choice of flocculation as a strategy to separate algae from culture media and provided a
proof-of-principle experiment for our planned genetic approach.

We repeated the flocculation efficiency experiment four times, varying the conditions and observing different results.
We also confirmed the occurrence of flocculation visually, by microscopy and a time lapse video (Figure 6).
When growth medium containing NaOH and CaCl2 was prepared in advance and added all at once at T=0,
flocculation and rapid sedimentation occurred almost immediately (Figure 6, 7) and cell clumping was clearly
visible under the microscope (Figure 8a-d). In contrast, gradual addition of base directly to cells followed by adding
CaCl2 at T=0 produced much slower and visually weak flocculation (Figure 9, 10a).
By switching out the wild type CC-125 for the starch over-accumulating strain dp120, flocculation was barely visible (Figure 10b).
This suggests that the mutant may respond differently to pH changes and metal cation addition compared to the wild type.

**Figure 6.** Timelapse video demonstrating natural sedimentation compared to chemically induced flocculation in
*C. reinhardtii*. Left: untreated culture showing natural sedimentation. Right: culture treated with NaOH and
CaCl2, where flocculated cells rapidly aggregate and settle significantly faster than untreated control.

![](https://static.igem.wiki/teams/5749/results/floc3.webp)

**Figure 7.** Chemically induced flocculation compared to natural sedimentation in *C. reinhardtii* ,
strains CC-125 and dp120. When the growth media was prepared in advance with NaOH and CaCl2, flocculated cultures
(red and blue) of both strains showed a much faster decrease in observed cell concentration compared to sedimentation
(green and purple), indicating accelerated settling when flocculation was induced.

![](https://static.igem.wiki/teams/5749/results/figure-3-ebc-results-cropt.avif)

**Figure 8.** Comparison of natural sedimentation and flocculation induced by addition of premade
TAP-media with added NaOH and CaCl2 in the cell wall deficient *C. reinhardtii* strain CC-3403.
**a.** Cells treated with NaOH and CaCl2 in 10x magnification.
**b.** Cells treated with NaOH and CaCl2 in 40x magnification.
**c.** Untreated cells in 10x magnification.
**d.** Untreated cells in 40x magnification.

![](https://static.igem.wiki/teams/5749/results/figure-4-ebc-results-cropt.avif)

**Figure 9.** Comparison of natural sedimentation and flocculation induced by addition of NaOH and CaCl2 directly to the
cells in the wild type *C. reinhardtii* strain CC-125. **a.** Cells treated with NaOH and CaCl2 in 40x magnification.
**b.** Untreated cells in 40x magnification.

![](https://static.igem.wiki/teams/5749/results/sed-floc-wt.webp)

**Figure 10a.** Chemically induced flocculation compared to natural sedimentation in *C. reinhardtii, CC-125*.
Cell concentration was measured over time for untreated, naturally sedimenting cells (blue) and cells treated with NaOH and CaCl2
to induce flocculation (red). For the wild type strain CC-125, linear regression analysis revealed a
significantly steeper decrease in cell concentration in the flocculation condition (p = 0.00762),
confirming that flocculated cells aggregated and settled faster than untreated controls.

![](https://static.igem.wiki/teams/5749/results/sed-floc-dp120.webp)

**Figure 10b.** Chemically induced flocculation compared to natural sedimentation in *C. reinhardtii, dp120*.
Cell concentration was measured over time for untreated, naturally sedimenting cells (blue) and cells treated with NaOH and CaCl2
to induce flocculation (red). Unlike the wild-type strain, no significant difference was observed between the two conditions for the starch
over-accumulating strain dp120 (p = 0.292).

## Electroporation

We optimized electroporation poring pulse voltages for *C. reinhardtii* using the linearized PLM005 empty vector in two
strains: the wild type CC-125 and the cell wall-deficient CC-3403.
Electroporations were performed with a 1 mm cuvette, applying two pulses for cell wall-intact cells and one pulse for
cell wall-deficient cells. In the first trial, CC-125 cells were electroporated with two poring pulses at 125 V.
No transformants were observed. In the second trial, CC-3403 cells were electroporated with a single poring pulse at 125 V.
Cell growth was observed for one of the two replicates (Figure 11). In the third trial, we tested additional voltages
(90 V, 105 V, and 120 V) on both CC-125 and CC-3403. No transformants were detected for either strain at these voltages.

Although plasmid integration could not be confirmed within the timeframe of the project,
these results suggest that 125 V may represent a suitable electroporation condition for the cell wall-deficient strain CC-3403.

![](https://static.igem.wiki/teams/5749/results/fig6-ebc.webp)

**Figure 11.** Electroporation result of cell wall-deficient *C. reinhardtii* strain CC-3403 at a
poring pulse of 125V, using a 1 mm cuvette. Transformed DNA is an empty vector of the PLM005 plasmid,
carrying paramomycin resistance. Growth was observed only on one of the replicates. Plates contain TAP agar supplemented
with paramomycin (20 µg/mL), carbenicillin (100 µg/mL) and L-Arginine 1:1000.

# **Summary**

During this project we decided on working with two different model organisms,
*E. coli*  and the green alga  *C. reinhardtii* . The aim was however the same, trying to express flocculation proteins
in the organisms. We were working with different protein types to see which would work better: FLO1 and Z17/Z18 pairs.
However, the idea was that these proteins would be expressed by the same mechanism, with a salt induced protomer.
Firstly, we tried making a cassette for the FLO1 protein, however, there were problems with ordering the gene, since it is
highly repetitive and we were unable to find a company that could synthesise the whole cassette of the salt promoter together with the
FLO1 protein. We therefore ordered the gene as three different parts that we would assemble with Gibson assembly,
however we did not have enough time to obtain the results. We therefore could not proceed with investigating its abilities to
flocculate  *C. reinhardtii* .

We also explored another set of proteins, the Z17/Z18 pair. We first started experimenting with transforming these proteins
into *E. coli* , which is a simpler organism to study expression and overexpression in. This cassette only contained the
proteins, without the salt protomer. We thereafter worked on isolating the protein pair, to study its interactions.
The proteins need a membrane anchor to be expressed on the surface of *E. coli* , but since we wanted to study the
purified protein interaction, and not cause flocculation of *E. coli* , we did not include an anchor in the cassette.
We managed to isolate and purify Z17 from the bacteria, however, this was unsuccessful for Z18.

Similarly, for Z17/Z18 to be expressed on the surface of the microalgae, it needed a membrane anchor, which is a problematic sequence
to synthesize when ordering a cassette. This anchor can be found naturally in the genome of  *C. reinhardtii*  and we therefore attempted to retrieve it by colony PCR, but this was unsuccessful. Despite these challenges, we were able to make
progress in other areas. Specifically, we optimized electroporation conditions in  *C. reinhardtii* , showing that poring
pulse voltages between 90–120 V were insufficient compared to 125 V in the cell wall-deficient strain CC-3403.
We additionally worked on developing an assay to evaluate flocculation efficiency compared to natural
sedimentation in  *C. reinhardtii* .

We also considered using another surface protein pair to induce flocculation - FUS1-MAR1. Unlike FLO1 and Z17-Z18, FUS1-MAR1 are endogenous to  *C. reinhardtii* , allowing us to clone the gene sequences from the genomic DNA of the algae by PCR,
circumventing the issues we were facing with getting the parts synthesised de novo. FUS-MAR1 are adhesion proteins,
which play a pivotal role in gamete fusion, however the pair has never before been used to induce flocculation in vegetative cells.
We planned to amplify the endogenous gene sequences and to place them under a strong promoter. Since FUS1-MAR1 functions as a
heterodimer, a population of algae, expressing either one of the proteins on its surface, would not flocculate together.
Unfortunately, we were not able to amplify the coding sequences for the pair, as cloning  *C. reinhardtii*
genes turned out to be much more complex than expected. However, through trial and error, we were able to optimize the protocol for cloning  *C. reinhardtii*  genes, which could be used by future iGEM teams.

In conclusion, even though the main goal could not be reached due to the lack of time, we succeeded with several important steps
along the way. We additionally worked on developing a method on how to measure flocculation, for future studies of flocculation
efficiency. For further research it would be interesting to continue our research and fulfill our goals of inserting the
different proteins into both  *C. reinhardtii*  to measure the flocculation, and to *E. coli* to study their
interaction more closely.

# **Outlook**

Our original aim was to transform constructed plasmids with flocculation genes (FLO1 respective Z17/18) into *C. reinhardtii*
and quantify their effect by comparing genetically engineered flocculation with natural sedimentation.
While we were unable to complete these steps within our timeframe of eight weeks, our experiments provided a
strong foundation for future work. We learned how to culture *C. reinhardtii*, demonstrated flocculation efficiency
through chemical induction, developed an assay method for comparing flocculation to sedimentation, and identified
electroporation conditions for the cell wall-deficient strain CC-3403 that warrant further exploration.

# **Appendix**

*#### pLM005 digestion*

We aimed to evaluate the compatibility of the BamHI and XbaI restriction sites when used in
combination. The PLM005 plasmid, which contains both recognition sequences, served as the template
for this analysis. Digestion was expected to generate four fragments, although one fragment is only
6 bp in length and was therefore unlikely to be visible on an agarose gel. The remaining fragments
were expected to appear as three distinct bands, see accompanying table.

**Table 1.** Amount of cuts and expected basepair length of pLM005 digested using BamHI and XbaI, single
enzyme and double enzyme digestion.

| Start | End | Length | Left Cutter | Left Overhang | Right Cutter | Right Overhang |
| --- | --- | --- | --- | --- | --- | --- |
| 1656 | 3023 | 1368 | BamHI | 5' | BamHI | 5' |
| 3024 | 3029 | 6 | BamHI | 5' | XbaI | 5' |
| 3030 | 4473 | 1444 | XbaI | 5' | XbaI | 5' |
| 4474 | 1655 | 4398 | XbaI | 5' | BamHI | 5' |

A total of four digestion reactions were performed:

* **Single-enzyme controls:** The PLM005 plasmid was digested separately with BamHI
  (Thermo Fisher Scientific) and XbaI (New England Biolabs, NEB). Each enzyme was used with its
  recommended buffer: FastDigest Buffer for BamHI (Thermo Fisher Scientific) and rCut Smart Buffer
  for XbaI (NEB).
* **Double digestions:** Two additional reactions included both BamHI and XbaI simultaneously.
  Each double digest was performed using one of the individual enzyme buffers (either FastDigest
  Buffer or rCut Smart Buffer) to evaluate buffer compatibility and enzyme efficiency under
  different conditions.

See Digestion protocol with the following modifications:

|  |  |
| --- | --- |
| DNA Input | 500 ng pLM005 (109.5 ng/µL concentration) |
| Reaction volume and reaction buffer amount | 50 µL total, 5 µL buffer |
| Restriction buffer | FastDigest (Thermo Fisher Scientific) and rCutSmart Buffer (NEB) |
| Restriction enzymes | FastDigest BamHI (Thermo Fisher Scientific), rCutSmart XbaI (NEB) |

![](https://static.igem.wiki/teams/5749/results/fig-6-bmc-results.webp)

**Figure 12.** 1% agarose gel electrophoresis of pLM005 digested with
restriction enzymes. Lanes: 1-GeneRuler 1 kb DNA Ladder; 2-XbaI and 3-BamHI,
single-enzyme digests; 4-Smart Buffer and 5-Fast Buffer, double digests with XbaI and
BamHI; 6-Water (pLM005), undigested plasmid control.

The successful digestions are represented in Figure 12. The biggest band is the undigested
supercoiled pLM005 plasmid (lane 6). Looking at the gel, all the expected bands were
obtained. What is more, to see is that the buffer composition did not affect the
functionality of the enzyme. Both bands (Figure 12, lane 4-5) are of same intensity,
implying that both buffers are as effective as the other.

*#### Z18 fragment and Z17-TEV-GFP-EhA plasmid digestion*

We aimed to digest the Z17-TEV-GFP-EhaA plasmid (hereafter referred to as pET plasmid) and the Z18
fragment with BamHI and XbaI to facilitate the replacement of Z17 with Z18.

The standard protocol recommended using 500 ng of plasmid DNA for efficient digestion. However,
500 ng of the Z18 fragment was not available. During previous restriction enzyme tests, 500 ng of the
PLM005 plasmid was successfully digested. To approximate equivalent molar amounts for the Z18 fragment,
we calculated the number of DNA copies using the following equation:

*copies = (ng \* (6.022 × 1023)) / (length × 109 × 650)*

Using this equation, the 500 ng of PLM005 plasmid corresponded to approximately 6.42 × 1010
DNA molecules. To achieve a similar number of molecules for the Z18 fragment, we calculated that
20.8 ng of the Z18 fragment would be required. This amount was used directly without purification in
the initial digestion.

A second digestion was performed using previously purified DNA (Z17-TEV-GFP-EhaA and Z18
fragments) (see Gibson Assembly protocol, step 2). Based on comparison with a DNA ladder, it was
determined that a 250 bp fragment required approximately 125 ng to be visible on a gel. Accordingly,
for a 25 µL digestion, 250 ng of the Z18 fragment was used to ensure adequate visualization of the
product.

See Digestion protocol with the following modifications:

|  |  |
| --- | --- |
| DNA Input | 500 ng Z17 plasmid (Z17-TEV-GFP-EhaA) (4 digests, each 50 µL reactions) and 250 ng / 20 ng Z18 fragment (2 digests, each 25 µL reaction) |
| Reaction volume and reaction buffer amount | 25 or 50 µL, buffer amount 2.5 µL or 5 µL respectively |
| Restriction buffer | FastDigest (Thermo Fisher Scientific) and rCutSmart Buffer (NEB) |
| Restriction enzymes | FastDigest BamHI (Thermo Fisher Scientific), rCutSmart XbaI (NEB) |
| Notes | Omit loading 20 ng Z18 fragment on gel |

![](https://static.igem.wiki/teams/5749/results/fig-7-bmc-results.webp)

**Figure 13.** 1% agarose gel electrophoresis of Z18 fragment and Z17-TEV-GFP-EhA plasmid, digested
with restriction enzymes. Lanes from left to right: (1) Thermo Fisher Scientific GeneRuler 1 kb DNA
Ladder; (2-3) Digested Z18 fragment with XbaI and BamHI; (4-15) Digested Z17-TEV-GFP-EhA
(pEt plasmid) with XbaI and BamHI; (16-20) Empty. Ran at 100V for 140 min.

After initial visualization under UV light, neither the DNA ladder nor the samples were
detectable. This observation was attributed to the accidental omission of SYBR Safe during gel
preparation. To remediate, 300 mL of 1X TAE buffer was combined with 30 µL SYBR Safe (10,000X),
and the gel was submerged in this solution. The gel was gently agitated at ~50 RPM and covered
with aluminum foil for 30 min to allow DNA staining. Total exposure of the gel and DNA to UV
light during imaging was approximately 1–2 min.

Figure 13 shows an analysis of Z17-TEV-GFP-EhA plasmid (pET plasmid) and Z18 fragment following
digestion with XbaI and BamHI. The digested samples were intended for subsequent excision and
purification using the Thermo Scientific GeneJET Gel Extraction Kit. Since Z18 is present only
as a DNA fragment, it is not possible to directly confirm a successful digestion as it was first
intended to create sticky ends on both sides. However, the plasmids in lanes 4–15 appear
linearized, as indicated by their uniform band size. This contrasts with the untreated pET
plasmid, which typically appears in multiple conformations, including supercoiled and circular
forms. This is then a good indicator of the successful digestion of our plasmid.

*#### Ligation troubleshooting of digested pET (TEV-GFP-EhaA) plasmid with Z18*

**NOTE: The subsequent transformation (see Transformation protocol) failed leading to troubleshooting
of the ligation protocol. See the results of a new digestion and ligation using pLM005 below.**

The Z18 DNA fragment was excised from an agarose gel and purified (see Gel Extraction Purification),
and the pET, TEV-GFP-EhaA, plasmid backbone was similarly gel-purified following digestion. In
parallel, an aliquot of digested Z18, that had not been run on a gel and therefore not undergone
gel purification, was retained as an additional ligation substrate (referred to as Z18-not-pure).

Followed ligation protocol, see BMC protocols with the following modifications stated in the table below:

| Z18-not-pure (µL) | Z18-pure (µL) | pET-plasmid pure (µL) | 10x-ligation buffer (µL) | T4-DNA-ligase (µL) | Replicates |
| --- | --- | --- | --- | --- | --- |
| 12.5 | 0 | 60 | 8.5 | 4.25 | 2 |
| 0 | 7.5 | 45 | 6.2 | 3 | 4 |

Additional modifications:

| Parameter | Details |
| --- | --- |
| Incubation temperature and time | Due to timing error, incubation time at room temperature was 25 minutes instead of 30 minutes |
| Downstream use/storage | Transformed into competent cells |

*#### Control - Digestion and Ligation using pLM005*

Due to the previously failed ligation of digested pET (Z17-TEV-GFP-EhaA) plasmid with Z18-fragment, a
control experiment of digestion and ligation enzymes was done. We decided to do a lot of controls by
changing a lot of parameters to try to assess the source of our previous failed experiments.

See Digestion protocol with the following modifications:

| Parameter | Details |
| --- | --- |
| DNA Input | 500 ng pLM005 (118 ng/μL) each in 5 tubes |
| Number of reactions | 5 mixes: XbaI, BamHI, double digest (XbaI + BamHI), control: no enzyme, control: ddH2O |
| Reaction volume | 50 μL |
| Restriction buffer type and volume | 5 µL rCutSmart buffer (NEB); ddH2O for water control |
| Restriction enzymes | XbaI, BamHI (single and double digests) |
| Controls | Two controls: (1) plasmid + buffer but no enzyme, (2) plasmid + ddH2O only |
| Downstream use | 6 µL |

See Ligation protocol with the following modifications:

| Parameter | Details |
| --- | --- |
| DNA Input | 6 µL (~60 ng) of digested pLM005 plasmid per tube, except for control containing non-digested pLM005. |
| Reaction design | 3 tubes with ligase (XbaI, BamHI, XbaI + BamHI digests), 3 identical controls without ligase, 1 control of non-digested pLM005 with restriction and ligase buffers but no ligase. |

**Table 2.** 1% agarose gel electrophoresis of digested and ligated pLM005 and controls. Numbers
representing lanes 1-20. GeneRuler 1kb DNA ladder from Thermo Fisher Scientific lanes 1 and 17.
Status of pLM005 plasmid (digested, ligated or control) and which digestion enzymes are used and
whether the samples are ligated are presented under corresponding lane.

| 1 | 2 | 3 | 4 | 5 | 6 | 7 | 8 | 9 | 10 | 11 | 12 | 13 | 14 | 15 | 16 | 17 | 18 | 19 | 20 |
| --- | --- | --- | --- | --- | --- | --- | --- | --- | --- | --- | --- | --- | --- | --- | --- | --- | --- | --- | --- |
| Ladder | pLM005 | Dig. pLM005 | Dig. pLM005 | Dig. pLM005 | pLM005 | - | Lig. pLM005 | Lig. pLM005 | Lig. pLM005 | - | pLM005 | pLM005 | pLM005 | pLM005 | - | Ladder | - | - | - |
| Thermo Scientific | ddH2O | XbaI | BamHI | XbaI, BamHI | - | - | XbaI | BamHI | XbaI, BamHI | - | XbaI | BamHI | XbaI, BamHI | - | - | Thermo Scientific | - | - | - |
| GeneRuler 1kb | - | - | - | - | Only restriction | - | T4 DNA ligase | T4 DNA ligase | T4 DNA ligase | - | - | - | - | Restriction buffer | - | GeneRuler 1kb | - | - | - |

![](https://static.igem.wiki/teams/5749/results/fig-8-bmc-results.webp)

**Figure 14.** 1% agarose gel electrophoresis of digested and ligated pLM005
and controls. Numbers representing lanes 1-20. 1-GeneRuler 1kb DNA ladder, 2-undigested
pLM005, 3-dig.pLM005 XbaI, 4-dig.pLM005 BamHI, 5-dig.pLM005 XbaI and BamHI, 6-control
pLM005 with restriction buffer without enzymes, 7-empty, 8-dig.pLM005 XbaI T4 DNA
ligase, 9-dig.pLM005 BamHI T4 DNA ligase, 9-dig.pLM005 XbaI and BamHI T4 DNA ligase,
11-empty, 12-dig.pLM005 XbaI, 13-dig.pLM005 BamHI, 14-dig.pLM005 XbaI and BamHI,
15-control pLM005 with restriction buffer and T4 DNA ligase buffer, 16-empty,
17-GeneRuler 1kb DNA ladder, 18 to 20-empty.

In Figure 14. we tried to see why previous ligation of pET (Z17-TEV-GFP-EhaA) plasmid failed.
To test the activity of the T4 DNA ligase (Thermo Scientific), ligation on pLM005 plasmid was
performed. At first, we loaded the digested pLM005 plasmid. This digested pLM005 plasmid was
then used for ligation.
However, it is good to point out that we had issues purifying the digested pLM005, which could
explain the faint bands in Figure 7. Another explanation could be due to a potential low
efficiency of T4 DNA ligase, which then could also explain the previous failed transformations.
This hypothetical decrease of efficiency could be explained by the bad storage of the enzyme
or by its age.
To conclude the failed transformation can be a result of multiple factors. Firstly, it was
challenging to obtain high concentrations of digested plasmids. Combining this with the lower
efficiency of T4 DNA ligase might have resulted in the unsuccessful transformations.

*#### Ligation and Transformation of Z18-pET plasmid*

***Z18 ligation***
Varying ratios of digested and gel-purified Z18 insert to the digested TEV-GFP-EhaA (pET) plasmid,
along with longer incubation times, were applied to the previously unsuccessful ligation protocol,
resulting in a successful ligation confirmed by transformation into competent cells.

See Ligation protocol with the following modifications:

|  |  |
| --- | --- |
| DNA Input | 6 µL (~42 ng) of digested pET plasmid per tube; insert (Z18) volumes adjusted to achieve 2:1, 3:1, and 5:1 insert-to-plasmid ratios based on molar calculations (see copy formula). |
| Incubation Time and Temperature | Overnight incubation for 11 hours at 22 °C. |
| Downstream Use | Transformation performed immediately after incubation. |

**Table 3.** The amount of copies of plasmid/insert relative to the length (bp) and total weight (ng)
presented. 4 calculations, consisting of three Z18-insert and one pET plasmid calculation for the
amount of copies needed to determine ratios, shown under sample type. Ratios are insert:plasmid.
Calculations determined using formula in ligation protocol.

| Sample | Length (bp) | Total Weight (ng) | Copies |
| --- | --- | --- | --- |
| Z18 5:1 | 300 | 9.265 | 2.86 × 1010 |
| Z18 3:1 | 300 | 5.559 | 1.72 × 1010 |
| Z18 2:1 | 300 | 3.706 | 1.14 × 1010 |
| pET Plasmid | 4986 | 42 | 7.80 × 109 |

***Transformation***
DH5α and JM109 competent cells were used for ligation of Z18 with TEV-GFP-EhA plasmid
(pEt plasmid) (see Transformation protocol for instructions).

**Table 4.** Transformation result of Z18-GFP-EhaA ligation (performed 30/7/2025). Ratios of Z18 to pEt
plasmid, spunn/centrifuged prior to plating, bacterial strain, and amount of colonies formed are
presented.

| Ratio | Spun (for plating) | Strain | Colonies |
| --- | --- | --- | --- |
| 2:1 | Yes | DH5α | 24 |
| 2:1 | No | DH5α | 1 |
| 3:1 | No | JM109 | 0 |
| 3:1 | Yes | JM109 | 0 |
| 5:1 | Yes | JM109 | 24 |
| 5:1 | No | JM109 | 1 |

{% endblock %}