Expression

After successful overexpression of the 3ɑ-HSD enzyme under pT7 promoter (BBa_K1674002), we conducted a series of experiments based on the 3ɑ-HSD activity measurement protocol, where we measured NADPH fluorescence over time added to E.coli lysates. Every experiment helped us understand a different aspect of the dihydrotestosterone (DHT) reduction reaction using our clones, and characterize the plasmid.

Enzymatic activity as a function of DHT concentration

The protocol for this assay can be seen here.

To get a basic idea of the kinetics of 3ɑ-HSD enzymatic reaction, we first wanted to examine the effect of increase in initial substrate concentration (DHT). Therefore, we sonicated E. coli BL21 cells containing BBa_K1674002 after two hours of induction with IPTG, added 150 µM NADPH to the lysates in a 96-well plate, and inserted into a plate reader pre-heated to 37℃ for 30 minutes to allow for result stabilization. 37℃ mimics human body temperature, which is the environment in which the enzyme will be working in our final product, and is in the optimal temperature range for the enzyme activity.(SOURCE)

In previous experiments we noticed fluctuations in fluorescence during the first 15-30 minutes after the addition of NADPH, even in negative controls, which we assume occurs due to a reaction of NADPH with the phosphate buffer, resulting in a new equilibrium state between NADPH and NADP⁺.

After 30 minutes we added DHT to each well in various concentrations and measured NADPH fluorescence during 5.5 hours. In a logarithmic time scale we can see linear behavior of NADPH degradation, where the slope represents the degradation rate (Figure 1). When comparing the different E. coli BL21 strains, with and without the 3ɑ-HSD gene, we can see clearly that the graph slope is steeper in presence of 3ɑ-HSD, implying faster NADPH degradation rate due to the specific enzymatic activity.

If we take the slope from each DHT concentration graph, we can describe the reaction rate as a function of the substrate concentration (Figure 2). It seems that the reaction rate in the absence of 3ɑ-HSD enzyme stays relatively constant with increasing DHT concentrations, whereas it rises logarithmically in the presence of the enzyme. From the results presented in Figure 2, we can assume that the enzymatic reaction rate reaches saturation at a DHT concentration between 40-60 µM. This result is compatible with the Michaelis-Menten enzyme kinetics model, which we described in detail in our modeling section.

Enzymatic activity as a function of NADPH concentration

The protocol for this assay can be seen here.

In the next experiment, we wanted to check the reaction rate dependence on the cofactor. Therefore, we performed the same procedure, this time with a constant DHT concentration of 50 µM, and varying NADPH concentrations. We observed a decrease in the reaction rate with an increase in NADPH concentration in presence of 3ɑ-HSD enzyme (Figure 3), contrary to substrate dependency. We were surprised by the difference between the effects of concentrations of substrate and cofactor, so we went back to our model to get some answers.

We hypothesize that during the 30 minutes of stabilization, some of the NADPH molecules were converted to NADP+ by other enzymes from the lysate which then could bind to 3ɑ-HSD and inhibits its activity. It is possible that in high concentration of NADPH, after 30 minutes, a large fraction of the 3ɑ-HSD molecules are bound to NADP⁺ and hence the reaction in the direction of DHT reduction is inhibited.

Enzymatic activity in lysates and supernatants of B.subtilis

Since our final goal is to express the 3ɑ-HSD enzyme in B.subtilis and to secrete it, our next step was to check the enzymatic activity in B.subtilis lysates and supernatants. In order to check for secretion, we designed a construct of the aprE signal peptide (SP) fused to the 3ɑ-HSD enzyme (as described in the secretion section).

In addition to the sonicated samples, we also took 1 ml from the supernatant of the B. subtilis bacterial cultures, which contains molecules secreted from B.subtilis during induction time. The reaction rate observed in the samples containing the signal peptide protein fusion was similar to that observed in the absence of the 3ɑ-HSD enzyme, indicating no enzymatic activity (Figure 4). It is possible the protein fusion damaged the enzyme active site, since we detected some activity in the lysates containing the 3ɑ-HSD enzyme itself, without the signal peptide. Further experiments are needed for verification of overexpression and activity, in order to optimize the conditions and achieve as much specific activity in supernatants as in E.coli lysates.

Secretion

Cofactor

Extracellular NADPH Assay

The protocol for this assay can be seen here.

We checked NADPH fluorescence at 340 nm excitation and 460 nm emission. We chose to check fluorescence as opposed to absorbance since it has been found to be more specific and sensitive method than absorbance. The NADPH measurements were done by reading fluorescence in a 96-well plate. The fluorescence readings were normalized by the culture’s O.D at 600 nm at a given time.

Glucose-6-phosphate dehydrogenase gene activity

At first, we wanted to see whether the zwf over-expression under the pT7 promoter ( BBa_1674004) in E.coli BL21 gave us the expected enhancement in NADPH. We observed that, as expected, the over-expression of glucose-6-phosphate dehydrogenase generated more NADPH, as can be seen in Figure 9, since there was a higher fluorescence/O.D._600nm in E.coli BL21 with zwf compared to E.coli BL21 wild-type.

Figure 9 illustrates that from time 0 until 12 hours, the fluorescence levels were similar in both in E.coli BL21 wild-type and in E.coli BL21 with zwf. After 12 hours a significant increase in the fluorescence/O.D._600nm was observed in the strain containing the zwf compared to the wild-type.

Consultations with academics led us to believe that NADPH is not excreted from the system. Therefore, we assume that the drastic change in fluorescence after 12 hours is probably because of natural lysis of the cells in the medium. If so, the values observed may give an indication of the amount of intracellular NADPH in the strains. This assumption is supported by the decrease in O.D._600nm parallel to the increase in fluorescence values at this point in time.

pgi and UdhA knockout activity

We performed the same experiment to check extracellular NADPH in culture with E.coli MG1655 wild-type and with the gene knockouts to see if the gene knockouts improved NADPH production. Additionally, we investigated the effect of adding a plasmid containing the zwf under the Rhl-R promoter and operon. For more information about the Rhl-R promoter and operon, see the cofactor project overview.

The results can be seen in Figure 10 below.

The results confirm that the E.coli MG1655 knockout has higher concentration of NADPH compared to the wild-type. However, the addition of zwf did not lead to the expected enhancement in NADPH. We hypothesize that after a long pefiod of time, the inducer of the RhlR promoter- C4HSL degraded and stopped the induction of the gene expression. This would explain the lack of difference between the extracellular fluorescence of E.coli MG1655 with the zwf gene and E.coli MG1655 without the gene. Another possible explanation is that the cells with the gene knockouts and zwf overexpression developed coping mechanism allowing them to partially restore metabolic equilibrium.

We concluded that the choice of an inducible, non-leaky promoter may not be efficient for our purposes. We plan create a clone with the gene under a constitutive promoter in order to better understand whether or not it is possible to get higher concentrations of NADPH with the addition of zwf into the E.coli MG1655 knockout.

Intracellular NADPH Assay

The protocol for this assay can be seen here.

In order to further investigate the effect of the over-expression of zwf with BBa_1674004, we measured the intracellular NADPH. In order to do so, we had to disrupt the cells without oxidizing the NADPH molecules. After a few unsuccessful measurements of the intracellular content using a sonication method, we decided to perform an experiment using various lysis procedures on E. coli BL21 with and without the zwf gene.

The results showed a higher intracellular NADPH concentration in the strain with the zwf gene for every lysis method used (Figure 11). Thus we concluded that the gene successfully enhances intracellular NADPH. The preferred lysis methods for NADPH concentrations, according to the results, are methods #3 and #7 (see protocol).

In the future, we hope to use these results to perform further experiments using the E. coli MG1655 knockout strain along with a plasmid for the overexpression of G6PD.

Integrative experiments

Commercial NADPH is both expensive and unstable. Therefore, to ensure sufficient amounts of the enzyme cofactor for the desirable reaction, our final product will include another bacterial strain, E.coli, which will be engineered to overproduce NADPH. We achieved this by enhancement of the zwf gene, which encodes for glucose-6-phosphate dehydrogenase (G6PD). This is a key enzyme in bacterial metabolism, since it is involved in the pentose phosphate pathway which is a main source of NADPH for the cell.

For the experiment, we took the extracellular medium of E.coli overexpressing the G6PD (BBa_K1674005) after 25 hours of growth, centrifuged it, and continued with the NADPH enriched supernatant, using the same protocol as for the activity check. The time at which the extracellular medium was taken was based on the results we observed, as presented in the cofactor results section of this page.

We performed the activity assay using 90 µl of this supernatant as the source of NADPH rather than commercial NADPH.

Comparing the reaction rate with 3ɑ-HSD enzyme and without it, we can see a minor difference (Figure 5), implying the specific enzymatic activity is lower than we have seen in previous experiments. We believe that the trend indicates that 3ɑ-HSD activity exists and that the NADPH supplied by overexpressed G6PD is adequate, yet further experiments are essential for isolating of the factors which might influence the enzyme environment.

Shelf-life