Difference between revisions of "Team:Austin UTexas/Results"

Kombucha Strains

The first steps in the characterization of microbes native to kombucha involved the isolation of strains from store-bought kombucha samples. This was accomplished by plating various dilutions of kombucha onto a variety of media including YPD, HS, and R2A.

Isolated colonies were selected from each "isolation plate" (Figure 1) and continually grown up and streaked out to ensure that the resulting frozen stock was truly axenic. Each newly isolated microbe was designated with a "KOM #" based on the order in which it was isolated (i.e. KOM 01, KOM 02, etc.) to serve as a placeholder name until the species could be identified. In order to begin this identification process, genomic DNA (gDNA) was first isolated from each individual strain. This DNA was then used as the template for two separate PCR reactions targeting either the 16S rRNA gene in bacteria, or the ITS rRNA gene for fungi. PCR products were then run on a 1% agarose gel to observe which reaction yielded product in gel (Figure 2).

Once it was determined whether each isolate was a bacterium or a fungus, the PCR products were purified and samples of the gDNA was sequenced using Sanger sequencing. The resulting sequences were then run through the Ribosomal Database Project (RDP) SeqMatch tool in order to identify the exact species of bacteria or yeast that correspond to each tested isolate. The identified microbes are listed below in Table 1.

Conjugation

We have attempted to conjugate GFP into both G. oxydans and G. hansenii with a Diaminopimelic Acid (DAP) auxotrophic strain of E. coli (The Barrick Lab). The plasmid contains the vector pMMB67EH, the promoter PA-1, GFP and a spectinomycin resistance gene.

The first conjugation was done with two of our isolated G. oxydans strains, in case the strains might behave differently. First, a mixture between our recipient strain and the DAP auxotroph strain were plated on an LB+DAP agar plate to allow for conjugation to occur. After 24 hours of incubation, we scraped up the growth and plated each conjugation mixture onto a LB+Spec plate (Figure 1). 24-48 hours later, we viewed the potential transconjugants using a fluorescence microscope. We then picked these glowing colonies and streaked them out onto another LB+Spec plate. We then followed our protocol for genome DNA isolation and 16S sequencing, as described above, to confirm successful conjugation of G. oxydans. After troubleshooting our 16s procedure, we were finally able to obtain a viable sequencing result. However, all of the glowing colonies were identified as E. coli.

For the next round of conjugation, we used a strain of both G. oxydans and Gluconacetobacter hansenii from the American Type Culture Collection (ATCC). Our procedure was identical to the one described in the previously paragraph.

We then picked isolated colonies and streaked them out onto LB+Spec plates. After using 16s sequencing on the potential transconjugants, we encountered an anomaly. Instead of amplifying the 16s gene, we received the sequence of the L,D Transpeptidase gene of E. coli.

We then decided to perform a Minimum Inhibitory Concentration (MIC) experiment in order to determine if G. oxydans is able to survive antibiotics above the standard E. coli concentration. We tested the antibiotics kanamycin, spectinomycin and carbenicillin.

Our results in Figure 2 showed that G. oxydans is resistant to at least 4x concentrations of spectinomycin (1x = 60µg/mL) and carbenicillin (1x = 100µg/mL). However, a 1x concentration of kanamycin (1x = 50µg/mL) was sufficient to inhibit growth. With this information, we then performed conjugations with E. coli donors that had a kanamycin resistance. These results are still pending through the wiki freeze.

Recapitulation

The process of recapitulations took individual microbes, isolated from kombucha, and recombined these to reform kombucha. This process is essential to create a designer beverage: the microbes that have been taken from kombucha and genetically engineered must be able to be put back together to remake kombucha. The process of recapitulation was repeated multiple times in triplicate. Each trial tested a different growing condition or different combinations of microbes for regenerating kombucha.

The first trials of recapitulations tested the tea media being used to compare the tea that at-home brewers used to see if this would support the microbial growth of isolates (Christensen, 2015). The next trials performed used strictly the microbes isolated from kombucha, the problem with this approach is that not all microbes that occur in kombucha can be isolated away from the symbiotic they exist in, and further they cannot grow on a solid agar plate (Imperial, 2014). Using the previous research on kombucha found in scientific literature, as well as our findings in lab, the most prominent microbes that help to create kombucha were attained (either by isolation from kombucha or by purchasing from a scientific database) to be used in future recapitulations (Dufresne, 1999 and Marsh, 2013).

After having the essential kombucha microbes, the next set of recapitulations tested what ratios these microbes needed to be in when placed in tea media so they could properly form a SCOBY and recreate kombucha. This process involved multiple trial and error results, but eventually the results included below were found.

Ethanol Reduction

Approach

Two potential ways to reduce ethanol content over the course of the fermentation process are to reduce the rate at which yeast produce ethanol or increase the rate at which bacteria convert the ethanol to acetic acid. The first of these methods is the most direct approach, and was the first method tested. We considered UV mutagenizing Lachancea fermentati, a yeast we isolated from kombucha, and then screening for ethanol production using a pH indicator, bromothymol blue (Figure 1). Bromothymol blue is blue at basic pH, turns green around pH 7, and turns yellow around pH 6, and has been used previously to screen for fermentation rate among Saccharomyces cerevisiae colonies (Robillard, 2007). During anaerobic respiration, both ethanol and CO2 are produced, and CO2 reacts with water to form carbonic acid, lowering the pH of the plate and changing the color of the indicator. A variety of problems with this approach led us to abandon it. It is likely that L. fermentati produce other acidic metabolic products, so pH would not necessarily correspond to amount of ethanol produced. This assay also relies on distinguishing differences in color in the agar to qualitatively determine amounts of ethanol production, a process that would be somewhat subjective. Additionally, the ethanol produced is necessary for the production of acetic acid, so slowing the rate of ethanol production would likely have slowed the fermentation of the beverage and could have thrown off the flavor. For all these reasons, attempting to decrease the rate of ethanol production by L. fermentati does not seem like a good approach to lowering the ethanol content during the fermentation.

We next considered increasing the rate at which acetic acid bacteria in kombucha convert ethanol to acetic acid. This would increase the amount of ethanol being used up throughout fermentation, ideally lowering the ethanol content. Two enzymes facilitate steps in this pathway (Mamlouk and Gullo, 2013). An alcohol dehydrogenase (PQQ-ADH) enzyme facilitates the conversion of ethanol to acetaldehyde, and a membrane-bound aldehyde dehydrogenase (ALDH) facilitates the conversion of acetaldehyde to acetic acid. In order to increase the rate at which ethanol is converted into acetic acid, we propose using Golden Gate Assembly to create a genetic construct in which expression of PQQ-ADH and ALDH is governed by a Tac-promoter (pTac), a hybrid promoter which is inhibited except in the presence of allolactose. The construct would be transformed into electrocompetent Escherichia coli and transferred to Gluconacetobacter hansenii via conjugation.

Identifying genes of interest

In order to design a construct increasing expression of PQQ-ADH and ALDH in Ga. hansenii, it was necessary to find the genome of the ATCC strain and identify the coding sequences for these genes. The whole genome shotgun sequence for our organism, ATCC 53582, is published on NCBI (Abbot, 2015) with annotations regarding the functions of specific sequences. Coding sequences are annotated with proposed gene products. Though there are several aldehyde dehydrogenase genes annotated in the genome, there is only one which is described as membrane-bound, matching the description from Mamlouk and Gullo (Mamlouk and Gullo, 2013). There are additionally multiple alcohol dehydrogenases. A known amino acid sequence for a homologous PQQ-ADH in Comamonas testosteroni was compared against sequences in the Ga. hansenii genome using BLAST (Table 1). One ADH enzyme found in the Ga. hansenii genome sequence matches the C. testosteroni sequence with a query cover value of 94% and an E value of 0 (third line of Table 1).

Creation of Golden Gate parts

In order to assemble the construct, the coding sequences for the genes of interest must be amplified from the Ga. hansenii genome and edited such that they have the correct Golden Gate overhangs and no internal BsaI or BsmBI restriction sites. The sequences were uploaded to Benchling for analysis and planning. The coding sequence for the membrane-bound ALDH contains a BsaI restriction site near the middle of the gene (Figure 2), and the PQQ-ADH coding sequence contains a BsmBI restriction site near the end of the gene (Figure 3). To eliminate the BsaI site in ALDH, primers were designed that would introduce a point mutation at the restriction site. One set of primers, igem2016_KOM_EtOH_01 and igem2016_KOM_EtOH_02, amplifies the sequence upstream of the restriction site, adding a type 3 Golden Gate prefix and removing the restriction site. Another set, igem2016_KOM_EtOH_03 and igem2016_KOM_EtOH_04, amplifies the region downstream of the restriction site, introducing a mutation to the site and adding a type 3 Golden Gate suffix to the end of the gene. These two products will be used in an overlap PCR reaction to create a final product with no BsaI restriction sites and the correct prefix and suffix for assembly. To remove the BsmBI site from the PQQ-ADH coding sequence, a set of primers (igem2016_KOM_EtOH_05 and igem2016_KOM_EtOH_06) was designed to amplify the region upstream of the restriction site and add a Golden Gate type 3 prefix to the beginning of the sequence. The reverse primer additionally adds a mutation to existing BsmBI restriction site and creates a new BsmBI restriction site that will be used to join the piece to a double-stranded DNA, igem2016_KOM_EtOH_07, containing the rest of the gene’s coding sequence appended with a Golden Gate type 3 suffix. The assembly of the PQQ-ADH part will therefore take place in two reactions: one reaction in which the upstream piece of DNA is created, and one reaction in which it is ligated to the gBlock. Table 2 contains more information about each of these oligonucleotides. All were ordered from IDT.

pH Sensors

During the kombucha brewing process, the beverage becomes more acidic. Additionally, it is unclear if or how the microbial community changes within the beverage over time. Thus, our team decided to find pH sensitive promoters that could be used to track not only the pH of the maturing beverage, but also the presence of the various microbes within the kombucha over time. We successfully created a neutral range reporter, attempted to create acidic and basic range reporters, and found three putative acidic range reporters that are endogenous to one of our kombucha bacteria, Gluconobacter oxydans

Though an acidic sensor was what was required for our kombucha analysis, the identification of sensors in other areas of the pH spectrum were explored as well. Three sequences were identified, the CadC operon for the acidic range, CpxA-CpxR complex for the neutral range, and the P-atp2 promoter from the BioBrick Registry (BBa_K1675021) for the basic range. Each sequence was paired with a unique corresponding reporter sequence so that if each pH sensitive plasmid were in the same environment, the specific pH of the system could be seen. The reporters used were, BBa_E1010 for the CadC construct, BBa_K1033916 for the CpxA-CpxR complex, and BBa_K592009 for the P-atp2 promoter.

CpxA-CpxR

CpxA-CpxR is a two-component mechanism that is activated at pH 7.4 and repressed at pH 6.0. CpxA is an intermembrane protein that autophosphorylates at a certain external pH, CpxR (a kinase) then gets phosphorylated by CpxA and acts as a transcription factor. This system originally is a transcription factor for the virF gene, but virF was replaced with a reporter. The original sequence was found in Shigella sonnei, but E. coli has a homolog of these proteins so all that is required on the construct is the appropriate prefix/suffix and CpxR binding site (Nakayama and Watanabe, 1995; Nakayama and Watanabe, 1998).

The order from left to right in figure 1 is control pH 6-9 and then Experimental pH 6-9. These are showing the gradient change in expression accordingly with the change of pH due to a pH-dependent promotor compared to consistent expression accordingly with a promoter that is always "on". The main point is that the control at pH 6 has more expression of the yellow-green chromoprotein than the Experimental at pH 6. The pH-dependent promoter of the experimental group is down-regulated at pH 6 whereas the control is not. Also, there is an increase in YGCP expression between the experiment pH 7 and pH 8 that is not seen in the control between pH 7 and pH 8. The normalized data in figure 3 shows the relative expression of YGCP. The CpxA-CpxR construct can be found on the iGEM registry as: BBa_K2097000, while the construct utilized as a control can be found on the iGEM registry as BBa_K2097002 as well as in figure 2.

CadC

The CadC operon is a native pathway in E. coli, involved in the cadaverine synthesis pathway. The protein CadC protein on the operon is produced and activates segments downstream of the operon on the CadBA receptors. The CadC protein is pH sensitive to an external pH 5.5 and below, as well as lysine dependent. A point mutation on codon 265, in which argenine is converted to cystine, causes the CadC protein to become lysine independent (Kuper and Jung, 2005).

Despite several attempts, we have been unable to grow the modified CadC operon on pSB1C3 in E. coli suggesting some form of cell toxicity. Specifically, all plasmids isolated have contained additional point mutations that appear to inactivate the expression of CadC. Due to this apparent toxicity, no data regarding the lysine-independent CadC could be collected. Alternative acidic range pH sensor candidates have been identified and are discussed below.

P-atp2

The P-atp2 promoter, native to the bacterium Corynebacterium glutamicum is reportedly induced at pH 7, to pH 9 (BIT-China-2015 and BBa_K1675021). Utilizing the blue chromoprotein (BBa_K592009), a test was designed in which a plasmid containing the P-atp2 promoter with the blue chromoprotein was grown alongside an E. coli line that contained a plasmid with just the blue chromoprotein. We expected to see constant blue chromoprotein production in the control series (those that lacked P-atp2) and a visual increase in blue chromoprotein as the pH was raised from 6 to 9 in the cells that contained the P-atp2 construct. The construct utilized as a control can be found on the iGEM registry BBa_K2097001 as as in figure 4.

However, as seen in figure 5, no clear change in color expression appears in the experimental trials, suggesting a lack of sensitivity of the P-atp2 promoter.