Team:Technion Israel/Project/Results

Team: Technion 2015



The goal of our project is to treat male pattern baldness by degrading dihydrotestosterone (DHT) on the scalp using synthetic biology tools. Throughout our project, we designed several parts with the aim of expressing a DHT reducing enzyme, 3α-hydroxysteroid dehydrogenase (3α-HSD), secreting it using a B.subtilis signal peptide, and producing enough NADPH as the reaction cofactor.

After addressing each level separately, we worked towards the assembly of the final design in a form of special 3D-printed comb which will combine the two types of bacteria in a combined friendly user experience. By testing the interactions between different parts, as well as the conditions required for bacterial storage, we were able to get as close as possible to accomplishing the vision of our new application.

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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.

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℃ is in the optimal temperature range for the enzyme activity.

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.

Figure 1 - NADPH degradation rate over time in logarithmic scale with initial concentration of 40 µM DHT

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.

Figure 2 - Reaction rate vs. DHT concentration

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.

Figure 3 - Reaction rate vs. NADPH concentration

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.

Figure 4 - Reaction rate in lysates and supernatants of different B.subtilis strains


After successfully cloning the mCherry reporter gene both alone and fused to the signal peptide from the aprE gene, we wanted to check whether the signal peptide actually works and secretes the mCherry to the growth medium.

mCherry secretion by B.subtilis PY79 wild-type

The protocol for this assay can be seen here.

In order to check whether the protein is secreted, we took two B.subtilis strains- one containing the gene for the mCherry protein and one with the signal peptide-mCherry fusion protein. After 2 hours of induction with IPTG 0.1 mM, samples were taken and centrifuged, and the fluorescence of the supernatant was measured at excitation wavelength of 587 nm and emission wavelength of 610 nm.

Figure 5 shows the fluorescence of the supernatant, normalized to O.D.600nm of each sample, as a function of time.

Figure 5 - Normalized fluorescence of supernatant vs. Time for B.subtilis expressing mCherry, with and without signal peptide

As can be seen in Figure 5, fluorescence of the supernatant of the strain containing only mCherry decreases over time, whereas the fluorescence of the supernatant of the strain containing mCherry fused to signal peptide initially showed an increase in fluorescence over time until reaching a maximum, and then followed by decrease over time.

The decrease in the fluorescence of the first strain is probably due to leftovers of fluorescent medium at the beginning of the experiment, and/or because of the cells which have undergone lysis, therefore causing mCherry to be present in the supernatant. The increase in fluorescence seen in the graph of the second strain suggests that the mCherry is secreted to the growth medium during the induction period, as expected. The decrease following that maxima is probably due to the instability of the protein in the extracellular environment and natural degradation.

It can also be seen that the fluorescence of the supernatant of the sample with the signal peptide is much higher than the supernatant of the sample with mCherry alone. In Figure 6, we examined the ratio of this difference by dividing the fluorescence values of the strain with the signal peptide by the strain without the signal peptide.

Figure 6 - Fluorescence ratio of strain with signal peptide-mCherry fusion to mCherry without signal peptide, over time

As can be seen in Figure 6, the fluorescence of mCherry secreted with the signal peptide is about 300% higher, relative to the fluorescence observed from the supernatant alone (with no signal peptide).

Since we observed degradation of the mCherry protein, we wanted to check out the differences between the degradation rates of the mCherry with and without the signal peptide.

Figure 7 represents a logarithmic time scale of the graph shown in Figure 5. The graph shows the degradation rates of the mCherry after reaching the fluorescence maxima.

Figure 7 - mCherry degradation rate (logarithmic time scale) after reaching fluorescence maxima

Figure 7 shows that the degradation rate of the secreted mCherry is 2.5 times higher than that shown by the not secreted mCherry, suggesting, again, that the mCherry is less stable outside the cell. Chemical reaction kinetics could explain this result, since degradation rate is mostly a function of mCherry concentration.


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_1674005) 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 8, since there was a higher fluorescence/O.D.600nm in E.coli BL21 with zwf compared to E.coli BL21 wild-type.

Figure 8 - Normalized Extracellular NADPH fluorescence vs. Time for E. coli BL21 strains

Figure 8 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 academic staff 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 RhlR promoter and operon. For more information about the RhlR promoter and operon, see the cofactor project overview.

The results can be seen in Figure 9 below.

Figure 9 - Normalized Extracellular NADPH fluorescence vs. Time for E. coli MG1655 strains

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.

In another experiment conducted in an identical fashion to the one mentioned above, we noticed peaks in extracellular fluorescence after 20 hours. We decided to check the ratio of the extracellular fluorescence for each of the strains, compared to E. coli MG1655 wild-type (Figure 10). The results reaffirmed the enhancement of NADPH production in the knockout strain, but also showed a slight increase in extracellular fluorescence in the knockout strain containing the zwf gene, when compared to the wild-type, unlike the results in Figure 9.

Figure 10 - Extracellular fluorescence in E. coli MG1655 strains compared to wild-type

The varying results can also be explained, as mentioned above, by inducer degradation or cell coping mechanisms.

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_K1674005, 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).

Figure 11 - Intracellular NADPH fluorescence in E. coli BL21 with and without the zwf gene

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.

NADPH Overproduction as a Function of Glucose Concentrations

We attempted to check the NADPH concentrations when the bacteria were grown with and without glucose in the medium. Glucose is a precursor of glucose-6-phosphate, the substrate of glucose-6-Phosphate dehydrogenase. Therefore, we wanted to check if the addition of more substrate would lead to more NADPH production.

This experiment’s results were not conclusive, so further experiments are needed.


For more accurate results in the future, we plan to use alternative methods such as HPLC, allowing the separation of NADPH from NADH, potentially giving us better insight into the NADPH concentrations in the clones.

Combined 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 12), 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.

Figure 12 - Reaction rate in E.coli lysate containing the G6PD enzyme


The protocol for this assay can be seen here.

Our goal in this experiment was to check the shelf-life and durability of our bacterial solutions containing glycerol and find the optimal conditions in which the product should be stored in stores and in the user’s home.

We used a solution of 80% glycerol and 20% LB (with or without glucose) in order to allow the viscosity of our solution to be such that the product won’t drip on the consumer's scalp.

Tubes mimicking the solutions that we expect to incorporate in the final product were prepared. We examined the durability and livespan of each bacteria used in this project, in the 80% glycerol, 20% LB solution, with or without the addition of glucose. The addition of glucose was in order to check whether or not it helps the survival of the bacteria in the different environments.

The samples were stored at various temperatures: 28˚C, 4˚C, and -18˚C, and the viable cells were check by live cell count, each day, for the duration of five days.


The results of the live cell counts for E. coli and B. subtilis over time can be seen in Tables 2 and 3 below. Figure 13 serves as a legend for understanding the symbols in the result charts shown in Figures 2 and 3.

Figure 13 - Legend: estimation of colony number for live cell count

The results of the live count from time zero are shown in Table 1.

The results of the live count for the different E.coli samples are shown in Table 2.

The results of the live count for the different Bacillus subtilis samples are shown in Table 3.


    Our results gave us insight into storage conditions which will be best for our product.

  • No significant differences in the number of colonies was observed with and without the addition of glucose, regardless of the storage temperature.
  • Bacteria which were stored at -18˚C resulted in many colonies after plating, over the course of the experiment. We also noticed that there was very little change in the consistency of the solution in this temperature, when compared to the other temperatures checked. This indicates that the consumer may be able to use the final product right out of the freezer with no need to defrost it first.
  • When stored at 28˚C, E. coli didn’t show growth in the live cell count after 24 hours. The B. subtilis samples, on the other hand, showed colony growth in all samples and growth conditions checked. However, the amount of colonies decreased from day to day, as shown in Table 3. For this reason, we should not recommend storing our product at room temperature.
  • When stored at 4˚C, the B. subtilis samples resulted in a significant amount of colonies. In the E. coli samples,however, the number of observed colonies decreased daily. Therefore, we should not recommend storing our product at 4˚C.
  • Based on the live cell count method, we should choose to recommend that users of our product store it in -18˚C.

Key conclusions

Our wide range of experiments resulted in several important findings:

  • 3ɑ-HSD enzyme activity in E.coli lysates is dependent on DHT and NADPH concentrations; whereas activity was observed to increase with increasing DHT concentrations, activity was observed to decrease with increasing NADPH.
  • The aprE signal peptide is sufficient to secrete the mCherry protein from B.subtilis.
  • Enhancement of the zwf gene in E.coli increases the production of intracellular NADPH, as well as extracellular NADPH over time.
  • NADPH supplied by overexpressed G6PD is adequate for the activity of 3ɑ-HSD.
  • The optimal condition for storing syringes containing the E.coli and the B.subtilis in an 80% glycerol medium is at -18℃.

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