The overall goal of this procedure is to demonstrate a system for generating unmarked mutants in the cyanobacterial Synechocystis species PCC6803 and marked mutants in Synechococcus species PCC7002. This method can help answer key questions in the cyanobacterial genetics field. Such as how to introduce chromosomal alterations into a strain.

By insertion of an antibiotic resistance gene, followed by a subsequent removal of this cassette, using a negative selectable marker. The main advantage of this technique is that strains can be repeatedly genetically manipulated. Allowing as many alternations to be introduced into a strain as desired.

After preparing media, cyanobacterial strains and plasmids, according to the text protocol. Set up a fresh culture by inoculating a loop full of cyanobacterial cells, into 30 to 50 mililiters of BG11 medium. Grow the culture in the light at 30 degrees Celsius for two to three days, to an OD7500, equal to 0.2 to 0.6.

Following the incubation, centrifuge one to two milliliters of the culture at 2, 300g's for five minutes. Then after discarding the supernatant, use BG11 medium to wash the pellets once, gently pipetting to resuspend the cells. As vortexing may result in the loss of pili, which are essential for DNA uptake.

After pelleting the cells again, and removing the supernatant, add BG11 medium to a final volume of 100 microliters, and transfer the cells to a 14 milliliter round bottom tube. Next, add 1 microgram of previous prepared plasmid A to the cells. And gently tap to mix.

Then lay the tubes down horizontally in the incubator at 30 degrees Celsius, and incubate for four to six hours. Following the incubation, spread aliquots of the cell culture plasmid DNA mixture on BG11 agar plates without antibiotics. And incubate at 30 degrees Celsius.

Approximately 24 hours later, prepare a 0.6%agar solution, in water containing kanamycin. And allow it to cool to 42 degress Celsius. Then add 2.5 to three milliliters to the edges of the culture plates and tilt the plate, so the solution forms an even top agar layer on the surface.

Incubate the plates approximately seven days for colonies to be visible. Then divide a fresh BG11 agar plus kanamycin plate into six sectors. And use a blunt end toothpick to streak out individual colonies.

To carry out PCR, to confirm marked knockout, transfer a small amount of cells into a tube containing 50 microliters of water, and approximately 20 425 to 600 micrometer glass beads. Shake the cells and beads at approximately 2, 000 rpm for five minutes, before spinning at 15, 700 g's for five minutes. Then use five microliters of the supernatant to prepare 50 microliter PCR reactions with tack DNA preliminaries.

To validate the mutants, after designing primers according to the text protocol, amplified product using the following program, and include a wild-type control. Through gelelektroforeses, verify the genotype of knockout transformants. Showing a band of approximately four kilobase pairs and lacking the wild-type band.

Refer to the text protocol for further details. If a wild-type band is still present, re-streak the strain on the fresh plate and repeat the PCR. Repeat the validation process until the mutant is segregated, and no wild-type band is observed in the PCR reaction.

For a strain that shows marked mutant profile, re-streak on a fresh plate for use in generating an unmarked knockout. To generate unmarked Synechocystis mutants, set up a fresh culture of the marked knockout by inoculating the loop full of cells into 30 to 50 milliliters of BG11 medium. Grow the culture for two to three days to and OD750, equal to 0.2 to 0.6.

Centrifuge 10 milliliters of the culture at 2, 300 g's for five minutes and discard the supernatant. Then us BG11 medium to gently wash the cells. After adding BG11 to a final volume of 200 microliters, and transferring the cells to a 14 milliliter round bottom tube, add 1 microgram of plasmid B DNA to the cells, and gently tap to mix.

Then lay the tubes down horizontally in the incubator, and incubate for four to six hours. Following the incubation, add 1.8 milliliters of BG11 medium and incubate the samples for a total of four days with shaking. This is a sufficient time to allow recombination to occur in the multiple chromosomal copies.

Plate 50, 10, and 1 microliter aliquots of the transformed culture onto BG11 plus 5%sucrose agar plates and incubate for approximately seven days to generate single colonies. Next, using a blunt end toothpick, hatch 30 to 50 individual colonies on BG11 plus kanamycin plates. Then patch onto BG11 plus 5%sucrose.

Colonies that grow on BG11 plus sucrose, but not on plates with kanamycin are potential unmarked knockouts. Colonies that grow on both plates are likely to be sucrose resistant due to a mutation in the sacB gene. Following the PCR method, demonstrated earlier in this video, verify unmarked knockouts, which will produce a gel band corresponding to the wild-type size minus the deleted region.

If the strain shows an unmarked mutant profile, then re-streak it on fresh BG11 agar without antibiotics. To store the strains long term, set up a fresh culture of the strain by inoculating a loop full of cells into 30 to 50 milliliters of BG11 medium. Grow the culture for three to four days to and OD750 equal to 0.4 to 0.7.

After using BG11 to wash the cells once, resuspend the pellet in approximately two milliliters of BG11. Add 0.8 milliliters of the concentrated cells to one tube. Then add 0.2 milliliters of 80%filter sterilized glycerol.

To another tube, add 0.93 milliliters of concentrated cells and 0.07 milliliters of DMSO. Store both tubes at negative 80 degrees Celsius. To revive the strains, remove the tube and use a blunt end toothpick to scrape off some cells onto an agar plate without antibiotics.

Then use a sterile loop to streak out as normal. This figure shows examples of plasmids A and B used to generate deletion mutants. In each case, the five prime and three prime planking regions are approximately 900 to 1, 000 base pairs.

Plasmid B, can also contain a gene cassette between the five prime and three prime approximately one kilobase pair flanking regions. Or a modified version of the native gene sequence. Upon transformation with plasmid A, several hundred colonies will appear on a plate after seven to 10 days.

If genes are non-essential and mutants demonstrate growth, similar to the wild-type strain, then all of the chromosomes should contain a copy npt1/sacB inserted sequence, as determined via PCR. If genes are non-essential, and mutants grow slow, then several rounds of re-streaking with increasing concentrations of kanamycin, are necessary to obtain a segregated marked mutant. If upon re-streaking, a marked mutant is not obtained, the gene is likely essential for survival.

The majority of individual colonies, obtained from transformation with plasmid B, will be kanamycin sensitive and sucrose resistant. As demonstrated here, PCR amplification of the target region in these colonies, shows that nearly 100%of the unmarked mutant profile. Once mastered, this technique can be done in hours over a six week period if it is performed properly.

While attempting this procedure it's important to remember to use sterile technique. Following this procedure, other methods like measuring oxygen production, or gas chromatography mass spectrometry, can be performed in order to answer additional questions, like measuring photosynthetic rates or metabolite production. After its development, this technique paved the way for researchers in the field of cyanobacterial genetics.

To explore key biochemical and physiological processes in this phylum, and developmental strains for industrial purposes. After watching this video, you should have a good understanding of how to generate unmarked knockouts in Synechococcus species PCC6803.