Team:Stony Brook/Experiments

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Experiments

Spectrophotometer

Step	Directions
1	Turn on spectrophotometer and wait for it to calibrate.
2	Click wave and set wavelength to 750 nm.
3	Fill cuvette with 1500 µL (three sets of 500 µL) of BG-11 medium. This will be the blank and will be placed in cell #1 (blue). Hit reference and the absorbance should equal 0.
4	Take culture from shaker or the ones at room temperature and spray them down with ethanol. Then, put them in the hood.
5	Spray the cuvette with ethanol and put them in the hood.
6	Using micropipette, fill the cuvette with 1500 µL of culture. Repeat with all cultures that need to be measured. You can use the micropipette to pipette up and down so that it’s homogenous.
7	Place cuvette in the cell, taking note of the cell number and which culture it came from. Make sure that the striated lines on the cuvette are facing towards the inside. The clear sides of the cuvette should be facing the light.
8	Close the lid, hit sample, and enter cell number.
9	Record absorbance number in the google sheets and wet lab notebook. Also, remember to include the time that the spectrophotometer measurements were taken.
10	After taking the measurements, discard the samples in the biological waste container.
11	Wash the cuvettes in RO water and flip them over to dry. DON’T FORGET to turn off the spectrophotometer at the end of the day or at the end of the use.

Nanodrop

Step	Directions
1	Click on Nanodrop app on computer (password for computer is on desk).
2	Select the material being sampled (nucleic acid or protein).
3	Blank nanodrop machine with water.
4	Use a micropipette to transfer 1-2 µL of sample onto machine.
5	Observe curve produced, take photo and document.

Plate Reader

Step	Directions
1	Check if anyone is using the plate reader → look at the start button, if in use, it would have a red stop button.
2	Open up software.
3	Insert desired settings in gear settings option.
4	Read area → can only select square are - can’t skip areas.
5	Press OK, plates should reflect settings on right hand column.
6	Must SAVE plates and data.
7	Put well plate into the plate reader.
8	If measuring absorbance, make sure that the correct filter is inserted.
8	Press read.

Cyanobacteria splitting/culturing

Step	Directions
1	Autoclave Erlenmeyer flask with cotton stoppers and aluminum caps (P1) (turn on autoclave 3 hours before use).
2	Follow hood procedure.
3	Warm up BG-11 media to 25 ℃ (work up to higher temperature 30 ℃).
4	Bring falcon tubes and autoclave flasks to hood (spray everything with ethanol).
5	Transfer desired volume of BG-11 to each flask using electronic pipette.
6	Transfer desired volume of cyanobacteria to flasks (mix in pipette)
7	Cover flasks, don't cross-contaminate.
8	Place flasks in shaker/incubator (start at 25 ℃, work up to 30 ℃).
9	Position lights in incubator.
10	Start shaker at 100 rpm.

Liquid inoculations: Agar to liquid

Step	Directions
1	Spray down the bench and hood. Spray the pipette and the plates that will be placed in the hood.
2	Set up the Bunsen burner.
3	Turn on the gas and use striker to light the flame.
4	Put 4 mL of growth media and antibiotics into the 15 mL culture tubes.
5	Disinfect the tungsten spreader by putting in in the flame of the Bunsen burner until it is bright orange.
6	Wait for the spreader to cool and test it by pressing it to the agar at the edge, if you don’t hear sizzling, it’s cooled.
7	Using the cooled spreader, pick up a colony and place it into the tube, swirl the spreader around to ensure that of all the colony is transferred.

PCR

Step	Directions
1	Set up and label 0.5 mL microfuge tube. Take out DNA fragment to be amplified, the appropriate primers, and Phire^TM Hot Start II PCR Master Mix from freezer to thaw.
2	Set up an ice bucket.
3	Mix the following in the 0.5 mL tube: Table 1 Recipe for general PCR reaction.
4	As the components are added to the tube, keep the tube on ice. This eliminates the slight chance that Phire^TM Hot Start II would activate at room temperature and catalyze DNA replication nonspecifically.
5	Start the appropriate PCR program on the thermocycler, wait until the temperature of the block is around 80 ℃, and then insert tube in.
6	Note: The PCR program is touchdown PCR, with the target annealing temperature a few degrees lower than calculated annealing temperature.
7	Note: for some constructs, the recipe was adjusted such that 1.5 µL of DNA and/or 2.0 µL of primer were used. The amount of water was also adjusted such that the total reaction volume was 50 µL.

PCR Purification

Step	Directions
1	Follow the protocol for the Monarch® PCR & DNA Cleanup Kit.
2	Derivations from protocol:
a	We performed an extra wash step.
b	When eluting, we added 10 µL of nuclease free water and allowed the DNA to incubate for 1 minute at 37 ℃. After eluting, we diluted the DNA by adding 5 more µL of nuclease free water.

Miniprep

Step	Directions
1	Follow the protocol for the PureYield™ Plasmid Miniprep System.
2	Deviations from protocol:
a	Before centrifuging the eluted solution, we left it at room temperature for 5 minutes.

Gel Electrophoresis

Step	Directions
1	Make 50 mL of 1x TAE solution in an Erlenmeyer flask, and pour in 0.6 g agarose. Do not swirl flask.
2	Microwave until solution is bubbling and homogenous.
3	Let the solution cool for 5 minutes and then add 1 µL of ethidium bromide. Swirl to mix.
4	Pour into gel casting tray, insert gel comb and wait for gel to set for 20-25 minutes.
5	Make 300 mL of 1x TAE running buffer in an Erlenmeyer flask. Add in 6 µL ethidium bromide and swirl to mix.
6	Put casting tray inside gel chamber and pour running buffer until gel is completely submersed. Carefully remove gel comb.
7	Load 15 µL DNA Ladder into one well. Load 20 µL of the DNA of interest mixed with 4 µL of 6x Gel Loading Dye into each of the other wells.
8	Note: When we used Diamond™ Nucleic Acid Dye instead of ethidium bromide to visualize DNA bands we followed the supplied protocol and made the gels without dye.

Cyanobacteria Transformation

Step	Directions
1	Prep plates with 10 ug/mL (high antibacterial concentration) of streptomycin.
2	Select a flask of cyanobacteria that has an optimal density of 0.6 to 1.0.
3	Take a 15 mL culture (contained in a falcon tube) and centrifuge for 10 minutes at 6,000 g.
4	Remove the supernatant and resuspend pellet in 10 mL of 10 mM NaCl.
5	Centrifuge again for 10 minutes at 6,000 g.
6	Remove the supernatant and resuspend pellet in 300 µL of BG-11 media and transfer to a microfuge tube.
7	Add 1-2 µL of desired DNA into the tubes and wrap tubes in aluminum foil to prevent exposure to light.
8	Incubate overnight at 30 ℃ with gentle agitation.
9	Plate cyanobacteria onto high concentration antibiotic plates. Wait 1-3 hours for plates to dry before placing into carbon dioxide incubator for overnight.
10	Pick a few single colonies to restreak (with an area of at least 1 square centimeter).
11	Make media composed of 46 mL BG-11, 9.2 µL streptomycin, 9.2 µL spectinomycin, and 460 µL of HEPES buffer.
12	Inoculate each restreaked colony in 2 mL of media and store in carbon dioxide incubator.

Ligation/Biobrick

Step	Directions
1	Digest 4 µL of the pSB1C3 linearized plasmid backbone provided in the iGEM Distribution Kit with 4 µL of the enzyme mastermix (at 37 °C for 30 minutes and heat killed it at 80 °C for 20 minutes). Table 2 Enzyme mastermix recipe for BioBricks

Workflow

Go with the flow.

Growth Experiments

After receiving our cyanobacteria cultures from Dr. Jackie Collier, we wanted to determine the optimal conditions for growth. First, we wanted to determine doubling time and growth rate at room temperature without a shaking incubator. We first made two replicates of a 1% solution of cyanobacteria (49 mL of BG-11 media and 0.5 mL of culture). In order to determine the density of the cultures, we utilized a spectrophotometer and measured the optimal density at 750 nm. We conducted measurements 3 times a day, spaced around 8 hours apart for 5 weeks. We used 750 nm to measure cell density because that wavelength does not conflict with that of the photosynthetic pigments such as chlorophyll.

Sodium bicarbonate experiments

For the second growth experiment, we wanted to determine the optimum amount of sodium bicarbonate to add to our cyanobacteria. The sodium bicarbonate acted as a buffer for the solution and as another source of carbon for the cells. First, we made a 1M filtered solution of sodium bicarbonate by adding 3.36 g of sodium bicarbonate powder to 40 ml of autoclaved Milli-Q®️ water and running the solution through a filter to filter sterilize the solution. We set up two sets of four flasks with a starting amount of 2% cyanobacteria (1 mL of culture in 49 mL of BG-11 media) for growth room temperature and 33 °C. The four different concentrations were 0 mM sodium bicarbonate, 5 mM sodium bicarbonate (250 µL of sodium bicarbonate solution) , 10 mM sodium bicarbonate (500 µL of sodium bicarbonate solution) and 20 mM sodium bicarbonate (1000 µL of sodium bicarbonate solution). We left one set of cultures at room temperature and grew the other set in a shaking incubator at 33 °C and 100 rpm. We conducted daily spectrophotometer readings for almost 200 hours.

Constructs/Plasmids

Plasmids were ordered from Addgene and arrived as agar stabs which we replated using a quadrant method. After overnight culture, we inoculated colonies from each plasmid in liquid Luria-Bertani (LB) media. After another overnight culture, we mini prepped using the PureYield™ Plasmid Miniprep System to isolate the plasmid DNA for use in NEBuilder®️ HiFi DNA Assembly with the constructs.

Constructs were ordered from Integrated DNA Technologies, Inc. (IDT) and their respective primers were ordered from Eurofins Scientific and they arrived in dry form in screw cap tubes. They then were resuspended with the appropriate amount of autoclaved Milli-Q®️ water based on the number of moles/weight of DNA shipped. Each construct was amplified via 50 µL PCR reaction. Reactions were set up as according to table 1.

All PCR reactions were done using the touchdown method, with an initial annealing temperature of 72 °C and gradually lowered to the calculated annealing temperature of the template DNA, usually in the mid to low sixties. Extension times were calculated based on template DNA length. The shortest fragments had calculated extension times of 3 seconds, which was inadequate. Thus, this was modified and we set the extension times for those fragments to 10 seconds. For some constructs the actual annealing temperature was set 3-4 °C lower than the calculated annealing temperature.

PCR cleanup was performed with the Monarch® PCR and DNA Cleanup Kit. We eluted with 10-15 µL of elution buffer or nuclease-free water.

To view DNA bands, we initially used Diamond™ Nucleic Acid Dye from Promega to dye our DNA. Dye solution was prepared by diluting the provided 10000x stock solution with 1x TAE. Gels were soaked in dye solution for around 30 minutes while being gently agitated on a shaker. The dye solution was most effective for a single use and was less effective in subsequent re-uses. When our supply of dye ran out, ethidium bromide was used in the running buffer and gel for visualization of DNA bands.

BioBrick Production

We digested the linearized plasmid backbone pSB1C3 supplied in the iGEM Distribution Kit with an enzyme mastermix (recipe shown in table 2), following iGEM Registry protocols with a few edits. Then, we ligated the digested backbone to purified PCR products of our constructs. We transformed the E. coli with the BioBricks following the iGEM Registry’s single tube transformation protocol and left the plated bacteria in an incubator at 33 °C for overnight. Next, we made liquid inoculations of the bacteria and grew them at 33 °C in a shaking incubator at 250 rpm for 12-14 hours. Afterwards, the vectors containing our Biobricks were isolated from transformed E. coli through miniprep. Successful transformation was confirmed by digesting the mini prepped products with restriction enzymes and running the digest on a gel. Furthermore, we sent in the BioBricks for sequencing for further confirmation of successful transformation. When we received our sequencing results, we isolated the BioBricks by miniprep, placed the solution of DNA on a 96 well microtiter plate and dried it out in a hood for 3 hours and then in an 60 ℃ incubator for 20 minutes to accelerate drying.

NEBuilder®️ HIFI DNA Assembly

In order to ligate our constructs together, we utilized the NEBuilder®️ HIFI DNA Assembly Cloning Kit from New England Biolabs. Following the protocol, we transformed DH5-alpha E. coli with purified PCR products. We left our plates in a 33 ℃ incubator overnight and the next morning, we made liquid inoculations of several colonies with LB media. These inoculations were grown in a shaking incubator at 33 ℃ and 250 rpm for 12-14 hours. Afterwards, we miniprepped our transformed E. coli to isolate DNA which we then used to transform cyanobacteria.

Cyanobacteria Transformation

The cyanobacteria were transformed using stock cultures of UTEX 2434 Synechococcus leopoliensis (from the UTEX Culture Collection of Algae). This culture is equivalent to Synechococcus elongatus PCC 7942 . We first conducted two experiments by plating the cyanobacteria on low (2 µg/mL) and high antibacterial concentration (10 µg/mL) of streptomycin and spectinomycin. We concluded that the high antibiotic concentration was more optimal for plating. We then proceeded to perform cyanobacteria transformation with plates made with the high concentration of antibiotics. When single colonies formed, we immediately restreaked a select few colonies onto a new plate (with colony patches greater than or approximate to 1cm2). After one round of re-streaking individual colonies were inoculated in BG-11 media with antibiotics and HEPES buffer.

Sucrose Assay Experimentation

In order to test CscB (sucrose permease), we added 150 mM of NaCl to 250 mL Erlenmeyer flasks with 50 mL of BG-11 to induce sugar production. S. elongatus PCC 7942 naturally produces sucrose to balance out the osmotic pressure caused by the extracellular salt. Additionally, we added 2 g/L of pH 8.0 HEPES buffer to prevent acidification of the media (as cscB depends on a basic environment to function). We also induced with 1 mM of IPTG after the cells had reached the mid-log phase.

The Sucrose/D-glucose Assay Kit was donated to us by Megazyme and was tested on the cyanobacteria after 1 week and then 2 weeks of IPTG induction. The cyanobacteria were spinned down at 10,000 g for ~5-10 minutes until a pellet formed, and 200 µL of their supernatant was extracted for the assay.

With the assay, the sample was mixed with both acetate buffer (negative control) and fructosidase. After incubating at 50 ℃ for 20 minutes, we added the GOPOD dye, which turns a bright pink (510 nm) when exposed to glucose. After incubating for the required time, we ran triplicates in a spectrophotometer and calculated glucose values with the calculator on Megazyme’s website.

We tested all of our samples against glucose and starch controls (0.25g/L of flour), as well as uninduced samples of cyanobacteria (which lacked NaCl and/or IPTG).

After the obtaining results, we measured the pH because CscB will not secrete sucrose in acidic conditions. Before even testing for sucrose, we noticed that the wild type culture (without antibiotics) had gotten visibly contaminated by fungus (figure 2). As such, we suspect contamination to be the major cause of our lack of results.

Then, we started a new round of testing, with and without added antibiotics. We conducted the sucrose assay as before, except the cultures with antibiotics were supplied with 10 µL spectinomycin and 10 µL streptomycin. The cultures we measured sucrose for are:

#1 = cscB #3 150 mM NaCl 1 mM IPTG 10 uM Spec 10 uM Strep 25 ℃

#2 = cscB #2 150 mM NaCl 1 mM IPTG 10 uM Spec 10 uM Strep 25 ℃

#3 = cscB #2 0 mM NaCl 1 mM IPTG 33 ℃

#4 = cscB #2 150 mM NaCl 1 mM IPTG 33 ℃

#5 = cscB #2 150 mM NaCl 33 ℃

#6 = cscB #4 #1 150 mM NaCl 1 mM IPTG 10 uM Spec 10 uM Strep 33 ℃

#7 = cscB #4 150 mM NaCl 1 mM IPTG 10 uM Spec 10 uM Strep 33 ℃

#8 = cscB #2 150 mM NaCl 1 mM IPTG 10 uM Spec 10 uM Strep 33 ℃

#9 = cscB #4 #2 150 mM NaCl 1 mM IPTG 33 ℃

Luciferase Promoters

In order to test the expression of our promoters, cpc, cpc560, idiA, psbA2, (which were incorporated into Dr. Susan Golden’s vector pAM1414 using Gibson assembly), we conducted luciferase experiments. Following Dr. Golden’s procedure, we added 5 µL of decanal to 95 µL of cyanobacteria in each well to induce expression. The decanal acted as a substrate for the bacterial luciferase enzyme, but due to the toxicity, the cells ended up dying, so the data obtained represents the end static expression. For the standard expression experiments for nighttime and daytime, we plated 95 µL of cyanobacteria into a 96 well plate, added 5 µL of decanal, parafilmed the edges and left the plate for 15 minutes before measuring the luminescence in a plate reader. For the high light experiment to psbA2 promoter, we plated 95 µL of cyanobacteria to half of the wells and put them under at least 500 µE of high light in our incubator for 1 hour. Then, we added 95 µL of cyanobacteria not exposed to high light to the other half of the wells and added 5 µL of decanal to all of the wells. Then, we placed the well plate in a plate reader and measured the luminescence. For the iron repressible experiment to idiA promoter, we plated 95 µL of cyanobacteria to half of the wells and exposed them to 2.3 mM of 2,2 dipyridyl (iron chelating agent) for one hour. Then, we added 95 µL of cyanobacteria not exposed to then iron chelating agent to the other half of the wells and added 5 µL of decanal to all of the wells. We placed the well plate in a plate reader and measured the luminescence.

2018 Stony Brook iGEM

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