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Revision as of 15:53, 15 October 2016
Opticoli
The new age of optics: Producing biological lenses and lasers to improve microscopy
Project Description
BIOLASERS
Imaging cells is essential for understanding life at the smallest scale and fighting cellular diseases like cancer. Imaging often relies on fluorescence, but fluorescent proteins have some drawbacks, such as their wide spectrum and low intensity.
Our biolasers will provide an accurate, safe and biological way to improve this.
Fluorescence is the ability of a molecule to take up the energy of a photon and release it again, which makes the molecule light up. Lasing works with the same principle as fluorescence, but now the light source is put between mirrors. The photons keep ‘bouncing’, increasing the energy of the system. When the light gets a certain power, the photons can escape in the form of a laser beam.
A biolaser is achieved by trapping fluorescent proteins inside a reflective agent. We have chosen two reflective agents: bioglass (polysilicate) and bioplastic (PHB). By covering a cell with polysilicate, the photons can resonate inside the cell, making a whole-cell laser. The polysilicate is synthesized by an enzyme called silicatein, which is expressed on the cell wall by fusion to membrane proteins. By filling a cell with PHB, which forms intracellular granules, the photons can resonate inside a part of the cell, making an intracellular laser. The PHB is synthesized after expressing the pha-operon. By fusing the GFP to the PHB synthase, the GFP is relocated into the PHB granules.
BIOLENSES
Microlenses are an emerging field in technology and have a ton of applications, including high-tech cameras, chips, solar panels and research & imaging techniques. However, they are expensive, hard to fabricate and the production uses heavy chemicals and high temperatures, so it is bad for the environment.
Our biological microlens will be cheap, easy to make and environmentally friendly.
When we cover a cell with polysilicate, using the enzyme silicatein, we are able to make a biological microlens. By overexpressing either the transcriptional regulator bolA or the cell division inhibitor sulA we can play with cell morphology and investigate optical properties. These enlarged cells can also be used in the lasing experiments. The single cell will be able to diffract light as a single microlens. When we make a grid of lenses, a microlens array, we can use the lens for a coating for solar panels, thin lightweight cameras with high resolution or 3D screens.
Experiments and results
Expression of different fluorophores
Introduction
One of the essential components of a laser is a fluorescent agent. Since our aim is to produce a fully biological laser, fluorescent proteins are favourable. To this end, we initially selected four fluorophores, with different emission wavelengths: GFP, mVenus, mKate, mCerulean. These fluorophores were reported to have an increased fluorescent intensity compared to their wildtype (Cormack et al., 1996; Nagai et al., 2002; Shcherboo et al., 2007; Rizzo et al., 2004).
Since mVenus, mKate and mCerulean did not exist in the iGEM registry yet, we constructed a brand new part for each of these, including strong constitutive promoter, RBS and terminators. GFP, on the other hand, was present in a whole range of biobricks. However, to our knowledge, there was no single biobrick available containing promoter, RBS and terminators. Hence, we constructed a new biobrick containing all of the above using the existing part E0840, consisting of RBS, coding sequence and terminators. By means of PCR we amplified this biobrick with primers designed to add a promoter while mainaining the biobrick prefix and suffix. Not only did we express GFP under the strong promoter J23100, but also under less strong promoters J23113, J23117, J23105, and J23108. This way we were able to see the influence of promoter strength on fluorescent output.
Experiments & Results
In order to characterize them before further use, the emission and absorption spectra were measured. Additionally, their effect on cell growth was investigated.
Emission spectra of different fluorophores
Introduction
To assure the fluorophores were functional, the emission spectra were recorded at the given excitation wavelength.
Methods
Parts encoding the four different fluorophores, including promoter, RBS and terminators, were expressed in E. coli BL21. GFP, mVenus, mKate and mCerulean were all expressed under the same strong constitutive promoter, J23100. Additionally, parts were constructed with GFP under control of promoters with different strengths, in order to investigate the influence of different fluorophore concentrations.
After growing in LB medium the cells were washed and resuspended in PBS of which aliquots of 100 µL were put in a 96 well plate.
The emission spectrum of each fluorophore was determined by exciting at a given wavelength and measuring the output intensity at a range of wavelengths. Because of this, the emission at a wavelength too close to the excitation could not be measured. This can be seen in the figures, where the left half of the emission peaks could not be measured. Especially for mVenus, where the excitation and emission wavelengths are very close together.
Results and Discussion
Fluorophores expressed under strong constitutive promoterAs all fluorophores were expressed under the strong constitutive promoter J23100, they were expected to show a strong fluorescence without the need of induction. Figure 1 shows that this was the case for GFP, mVenus and mCerulean. mKate, however, did not show any fluorescent activity and was therefore not used in the subsequent steps of the project.
Not only did we express GFP under the strong promoter J23100, but also under less strong promoters J23113, J23117, J23105, and J23108.
For comparison, the emission was normalized by dividing by OD. All strains were measured in the same dilution, in order to make the results reproducible. The emission intensity is as expected, according to the promoter strength. All fluorophore spectra were also recorded in a dilution more suited for their emission intensity and normalized by 1 (Figure 3). From this figure we can conclude that all GFP biobricks function properly.
The effect of expression of fluorophores on growth
Introduction
As consitutive expression can sometimes be hard on the cells, we investigated the effect of the different consitutive promoters on cell growth. In a 24 hour kinetic cycle alterating shaking at 37°C with fluorescence and optical density measurements, we investigated whether this was the case.
Methods
An overnight culture in eM9 medium was inoculated in fresh eM9 to an OD of 0.1 in a 96 well plate. The emission at 522 nm was measured every 15 minutes. Measurements were done in quadruplicate with pure eM9 as a blank.
Results and Discussion
Figure 1 shows the 24 hour measurement of optical density and fluorescence intensity. The final OD is approximately equal for all different strains, suggesting that the level of constitutive expression was not influencing the growth. Furthermore, fluorescence intensity drops after the exponential growth phase, suggesting that GFP is being broken down by proteases as a response to nutrient limitation. After this event, growth continues at a slower pace, while GFP activity keeps decreasing. All in all, constitutive expression of GFP does not seem to have a detrimental effect on cell growth during exponential phase.
Cells expressing mCerulean or mVenus, however, seem to be having a longer lag phase before exponential growth starts (Figure 2). Also, they reach a lower final OD than the ones expressing GFP. These fluorophores might be slightly harmfull for cell growth. Nonetheless, they do grow and exhibit fluorescence, so they can be used in further experiments.
Discussion & Conclusions
We were able to succesfully construct and characterize two biobricks with brand new fluorophores for the iGEM registry: mVenus and mCerulean. Also, we constructed five new composite parts, based on the existing GFP biobrick. All in all, these new parts provide a ready-to-go expression device for green, yellow, or cyan fluorescence, which can be very usefull for future iGEM teams.
Coating the cell in polysilica using silicatein
Introduction
For both the biolaser and the biolenses we need a coating of polysilicate, biological glass, around the cell. For the biolaser this glass will form the cavity that will enable the cells to emit laser-like light. For the biolens, the glass will give optical properties for the cell. E. coli is intrinsically not able to coat itself in polysilicate. However, upon transformation of the silicatein-α gene, originating from sponges, it is possible to coat the bacterium in a layer of polysilicate (Müller et al., 2008; Müller et al. 2003). Therefore, we are transforming E. coli with silicatein-α. We test the use of two different silicateins, one originating from the marine sponge Suberites domuncula (Müller, 2011) and one originating from the marine sponge Tethya aurantia (Cha et al., 1999). We express the enzyme in three different ways. First of all, we expressed the gene from S. domuncula and see if the enzyme is transported outside the cell as described by Müller et al, 2008. Furthermore, we express a fusion of silicatein from T. aurantia to the trans-membrane protein OmpA (outer membrane protein A) from E. coli to anchor the silicatein to the membrane (Curnow, Kisailus, & Morse, 2006; Francisco et al. 1992), which might make coating the cell specifically in polysilicate more efficient. We also express a fusion of silicatein from S. domuncula to the transmembrane Ice Nucleation Protein (INP) from Pseudomonas syringae, a popular protein for membrane fusions (Kim & Yoo, 1998). Using these different approaches we expect to coat the cell in polysilicate, an overview is shown in figure 1. This glass coating around the cell will be the basis of both our biolens and –laser.
Experiments & Results
To determine if we have successfully covered E. coli in polysilicate, and to characterize the properties of the silicate-coated cells, we have performed a series of tests. First of all, we have stained the cells with rhodamine, a fluorescent stain that is able to bind to polysilicate. These stained cells were observed under a fluorescence microscop to determine whether the polysilicate shell was present. Furthermore, the polysilicate-synthesizing cells were observed using both Scanning Electron Microscopy (SEM) and Transmission Electron Microscopy (TEM). We have also determined the physical properties using Atomic Force Microscopy (AFM) to see whether the polysilicate layer changes the stiffness of the cells. Lastly, we have also performed a growth study of the polysilicate-coated cells to determine whether the polysilicate layer affects growth of the organisms.
Rhodamine staining of silica covered cells
Introduction
After transforming E. coli with the different silicatein BioBricks, we wanted to confirm and image whether the cell had indeed synthesized a layer of polysilicate around itself. Before using any advanced imaging techniques, we first used a much simpler technique. It is possible to stain polysilicate depositions with the fluorescent stain Rhodamine 123. This stain has shown to bind specifically to polysilicate (Li, Chu, & Lee, 1989). Since the stain is fluorescent, we can image it with a simple fluorescence microscope. If a cell has a polysilicate layer, the Rhodamine will bind to it which we can image.
Methods
The experiment was performed using E. coli BL21 with the following plasmids and conditions. All genes are under an inducible promoter.
| Plasmid(s) | IPTG | Silicic acid | Rhodamine 123 |
|---|---|---|---|
| OmpA-Silicatein | + | + | + |
| Silicatein | + | + | + |
| INP-Silicatein | + | + | + |
| OmpA-Silicatein (negative control) | + | - | + |
| OmpA-Silicatein | + | + | - |
The cells were stained with 0.1 vol% Rhodamine. And washed 5 times with PBS, prior to imaging (Li et al., 1989; Müller et al., 2005). Both widefield- (light microscopy) and fluorescence microscopy were used to image the cells.
Results and discussion
The stained cells were first imaged at maximum excitation intensity. At this excitation energy, only OmpA-silicatein showed fluorescence specifically localized at the cells and not in the medium. Silicatein, INP-silicatein and the negative control all caused overexposure of the camera. In figure 1 the imaging results of OmpA and the negative control are shown. The cells that were not stained with Rhodamine showed no measurable fluorescence. (data not shown).
Since Silicatein, INP-silicatein and the negative control all caused overexposure of the camera, they all had the same output. We can thus not draw any conclusions for these strains. Therefore, samples where overexposure was observed were imaged again at only 1/3 of the excitation energy. The imaging results are displayed in figure 2.
At this excitation energy, we can compare these samples. From figures 1 and 2 we can see that the strain transformed with OmpA-silicatein clearly has a different output from the negative control. The fluorescence is only localized at the cells. From this we can conclude the Rhodamine has stained the cells and therefore these cells will contain the polysilicate layer. We cannot distinguish a clear difference between silicatein, INP-silicatein and the negative control. The entire medium is fluorescent, which causes overexposure of the camera at high excitation intensity. This might mean that the Rhodamine is not specifically located at the cell walls, but still dissolved in the medium. Wedo see some fluorescence localized at the cells, but the diference between the fluorescence of the medium and the cells is much smaller than we observed for OmpA-Silicatein. Therefore, we cannot conclude that the strains transformed with silicatein and INP-silicatein are able to synthesize a polysilicate layer around the cell. We might be able to test this using SEM or TEM, but from this test we can not draw a conclusion for these two strains.
Back to TopImaging of silicatein-expressing cells using SEM
Place experiment (Introduction, methods, results&Discussion here)
Back to TopImaging of silicatein-expressing cells using TEM
Place experiment (Introduction, methods, results&Discussion here)
Back to TopAnalysis of physical properties of silica covered cells using AFM
Place experiment (Introduction, methods, results&Discussion here)
Back to TopViability experiments of silica covered cells
Introduction
Since the silicatein expressing cells are to cover themselves in polysilicate, their nutrient supply might be limited by diffusion, which can eventually result in cell death. To investigate whether this is indeed the case,a growth study was performed.
Methods
Cells containing the three different silicatein biobricks were grown overnight in selective LB. They were transfered to fresh medium and grown until in exponential phase. Then IPTG was added to induce expression. After a subsequent incubation of three hours, the medium was supplemented with silicic acid as substrate for silicatein. During the following five hours samples were taken, of which a 10-6 dilution was plated on selective LB plates. Colony forming units (cfu) were counted the day after.
Results and Discussion
Cells expressing either silicatein from S. domuncula (Sil Sdom), silicatein from S. domuncula fused to INP (INP Sil Sdom) or silicatein from T. aurantia fused to OmpA (OmpA Sil Taur) were tested. As a negative control, OmpA Sil Taur expressing cells without silicic acid were used. After one hour no colonies were observed on the plates on which the cultures with silicic acid were plated (Figure 1). The cultures without silicic acid continued to grow until after five hours.
Discussion & Conclusions
Conclusion and discussion on the experiments
Engineering a biological laser
Introduction
Our cellular laser consists out of two features: fluorophores will be the light of the laser and a silica layer synthesized by silicatein will be the ‘mirrors’ that reflect a part of the photons emitted by these fluorophores. The fluorophores first need to be excited by an external light source. This could be either an LED source or a conventional solid laser. For fluorescence an LED excitation source suffices. However, we want lasing to happen in our cells. For this to happen, we need ‘population inversion’, which means the majority of the fluorophores is in an excited state (Gather & Yun, 2011; Svelto & Hanna, 1976). More information on this can be found in the project description. In order to excite the majority of the fluorescent proteins at the same time, we need a strong excitation source. Therefore we need to use a laser to excite the fluorphores in our biolaser. However, a major downside to using lasers for fluorophore excitation is the occurrence of photobleaching (Eggeling, Widengren, Rigler, & Seidel, 1998). The laser power required for the excitation of fluorophores to induce population inversion in the cell is so high it will photobleach the fluorophores within microseconds (Jonáš et al., 2014). Therefore, we need a custom laser setup to prevent photobleaching but establish the population inversion required for our biolaser. This laser setup should contain a pulsing laser which pulses at a frequency that will maintain the excited state but does not photobleach the proteins.
The appropriate set-up was not available, so we decided to build our own microscope out of separate optical parts. By discussing our problem with optics- and photonics companies and showing our motivation to solve this challenge, we were able to get all our required components sponsored or borrowed, making the entire set-up nearly cost-free. With our minimal set of available tools, we calculated and designed the optics in such a way that we could image fluorescent cells, while photobleaching was minimized. After days of laser aligning, we managed to do so. More information on the design of this setup can be found on the hardware page.
Using our custom-built setup, we analysed our ‘Biolaser’-cells, to see if they were able to produce a laser-like emission of light.
Experiments & Results
The first and foremost experiment to be done with the setup was to image the cells in order to see if the setup works properly and to see whether we can image cells with it. Once we have confirmed that the setup works properly, we can measure the output intensity of the cells to see whether the cells are able to emit laser-like light.
Imaging our biolaser using our home-built setup
Introduction
Building an optical setup is a very precise work. First, it is important to calculate the positions of all components in such a way that the laser beam will reach your sample, and the light emitted by the sample consequently reaches the detectors of the camera and spectrometer. Once this is done, all optical components are positioned on an optical table. This special table is made to prevent vibrations in you system and has mounting holes so all optical components can be screwed into place. Once the components are positioned on the table, the alignment begins. In this step, the beam coming from the laser is guided throughout the system. By slightly adjusting and repositioning all optical components the light is guided through the components until it reaches the detector of the camera. It is essential that the components are aligned correctly and are free of vibrations, because this could change the path of the light.
After tens of hours of carefully placing components and aligning the light through the setup, we managed to direct the light from the laser, through all components onto the detector of the camera. However, this does not necessarily mean that when we add fluorescent cells to the setup it we are able to image the cells with the setup. Any error in the setup could cause it not to work. Therefore, we first had to confirm whether our setup was working.
Methods
In order to confirm whether the setup was working, we used E. coli BL21 cells that were transformed with our constitutive mCerulean BioBrick. We have previously confirmed that these cells are able to fluoresce and that they can be excited at 405nm, the wavelength of our laser. To make sure the only output we were measuring was fluorescence, and not any ‘leakage’ of light from our laser beam, we also tested cells that were not transformed with the mCerulean BioBrick. These cells are not able to fluoresce after excitation at 405nm, so if the setup is working properly we should not get a signal from these cells.
The cells were fixated to the microscope slides using 3% agarose pads and imaged at an excitation intensity of 0.5 mW. This energy is low enough to not instantly photobleach the proteins, but observe fluorescence clearly. Focusing on the cells was done manually with a 50x oil-immersed objective. The cells were excited with a Coherent OBIS LX 405nm laser and images were taken using a DeltaPix Invenio III CCD camera.
Results & discussion
Imaging the cells with the setup yielded the following results:
As we can see in figure 2B, the cells transformed with a gene for the fluorescent protein mCerulean are clearly visible. When using a strain transformed with a plasmid that is not known to cause the cells to fluoresce, in this case OmpA-silicatein, no light was observed, as shown in figure 2A. From this we can conclude that we successfully built a setup that is able to observe and measure fluorescence in a cell. There is no leakage of light of our excitation laser in the camera, since we do not observe anything when we use non-fluorescent cells. Also at a higher excitation energy (50 mW) we did not observe anything on the camera. Therefore we can conclude that our setup works as expected as it is indeed able to measure fluorescence without measuring other light sources.
For this experiment we used E. coli transformed with OmpA-silicatein that was induced and incubated in silicic acid, so it would contain the silica layer. We used this strain both as a negative control as well as to test whether the silica or the cells had any autofluorescence that could interfere with our laser experiments. We did not observe any fluorescent signal for these cells, so we can conclude that the silica layer does not have any autofluorescence at 405 nm. Therefore, these cells are suitable for the laser experiments.
Back to TopIntensity measurements of laser cells
Introduction
One of the biggest differences between a laser and fluorescence is the amount of emitted photons. We can measure the amount of emitted photons by measuring the intensity of emitted light. The intensity of fluorescence increases linearly with the excitation energy. However, at a certain excitation energy it will reach the so-called ‘laser threshold’ and stimulated emission, and thus lasing will occur. From this point on, the output intensity of the fluorophores will still increase linearly, but with a much steeper slope, as shown in figure 1.
We investigated whether we could find the same relation between input and output intensity for our Biolaser cells, using our self-built setup, to investigate whether our cells were able
Methods
For this experiment we used E. coli BL21 transformed with the following BioBricks:
| Plasmid(s) and conditions | Function |
|---|---|
| mCerulean (constitutively expressed) | Fluorescence |
| mCerulean (constitutive) + OmpA-silicatein (induced, incubated in silicic acid) | Biolaser |
| OmpA-silicatein (induced, incubated in silicic acid) | Negative control (no fluorescence) |
The cells were fixated on a microscope slide using 3% agarose pads. The cells were excited at a wavelength of 405 nm. The cells were imaged at excitation energies of 0.1 mW, 0.5 mW, 0.7 mW, 1 mW, 2 mW, 5 mW, 10 mW and 50 mW. These images were analysed using ImageJ to determine the output intensity and corrected for background noise (McCloy et al., 2014).
Results and discussion
The output intensities of our cells were plotted against the excitation power to determine whether our cells emitted laser-like light. The results are shown in figure 2.
In figure 2, the measured intensity is plotted against the excitation power (black dots). Since we expect a linear relation, the expected increase of fluorescence was also plotted (in a blue line). If the slope of the measurement data is higher than this slope, we observe lasing. If the slope is lower than the expected line, we observe photobleaching. In figure 2 we see that the measurement data first nicely matches the expected increase in fluorescence. However, at an excitation power of 2 mW we see that in all cases the fluorescence stays at the same level. This means that the intensity is not increasing anymore and we observe photobleaching. So, both the fluorescent cells as well as the laser cells do not emit any laser-like light. For the fluorescent cells that were transformed with solely mCerulean, this result is as expected, these cells do not have any extra modifications that should cause them to emit laser light. The strain that was transformed with both mCerulean and OmpA-silicatein had a glass shell around the cell that could cause the cells to emit laser light. This effect was not observed. The strain transformed with solely OmpA-silicatein did not emit any light (results not shown).
From this experiment we can conclude that the cells were not able to emit laser light. This could be both due tour setup or due to the constructed cells. The lasing should have occurred at an excitation power under 1mW (Fan & Yun, 2014). However, up to this excitation power the Biolaser cells follow the same slope as the fluorescent cells, which means that no lasing occurred. From 2 mW and higher the cells bleach. Since we expected the cells to lase at an excitation energy under 1 mW, we did not take such high energies into account while designing our setup. Even though we exposed the cells to the excitation laser for a very short time, the power was so high that the cells eventually photobleached anyway.
Even though the excitation laser eventually bleached the fluorophores, this was probably not the reason why lasing didn’t work. Lasing should have occurred at an excitation energy between 0 and 1 mW, and at these energies the fluorophores did not bleach. This indicated that there could be another reason why the cells did not lase. Modeling showed, that in a cavity as big as E. coli (around 1 µm) we need an intracellular fluorophore concentration of 0.1 M to get lasing. In our cells the maximum concentration we can achieve is in the nM to µM range and therefore lasing physically not possible in our cells. To get lasing, we would either need much larger cells or a much higher concentration of fluorophores. Therefore, it was most likely not due to the self-built setup that we did not observe lasing.
Back to TopDiscussion & Conclusions
We have successfully calculated, designed and built a custom optical setup that could allow us to measure lasing in cells. Using this self-built setup, we were able to image fluorescent cells and measure the intensity of the fluorescence. We have confirmed that the setup worked. However, we did not observe any lasing in cells. Our models have shown that this is due to the size of our cells, if we would use a bigger organism, e.g. a mammalian cell line, we could be able to measure lasing. Unfortunately, we were not able to measure this in our setup, since we did not have the safety permit to use mammalian cells. However, this would be a very interesting future study.
Engineering biological lenses
Introduction
Introduction on lenses & experiments
Experiments & Results
Introduction on the experiments that we did
Influencing cell shape for round lenses
Introduction & background
When making biological lenses, the shape of the lens is of crucial importance. E. coli is a rod-shaped organism, so it’s not symmetrical along all axes. Shining light on the round parts of E. coli has a different effect on the focusing of light than shining light on the long sides, see figure 1. More information on this can be found on the modeling page.
For some applications, such as the solar cells, this variation in shape does not matter that much; here it’s most important that light gets focused in any way. However, when we want to use our microlenses in more advanced optical systems, such as microscopes or cameras, we need to make sure that this variation between the different lenses is minimized. Manufacturers of optical systems do not accept a high aberration between different lenses, so it’s crucial for us to be able to control the shape of our lenses. We have decided to engineer E. coli in such a way that it becomes spherical. This way we are able to create spherical lenses. Apart from the fact that it is crucial to be able to control cell shape, round cells offer the advantage of being symmetrical along all axes, so the orientation of your lens does not matter for the optical properties.
In order to create spherical E. coli, we overexpress the BolA gene. BolA is a gene that controls the morphology of E. coli in the stress response (Santos, Freire, Vicente, & Arraiano, 1999). By overexpressing this gene, the rod-shaped E. coli cells will become round (Aldea, Hernandez-Chico, De La Campa, Kushner, & Vicente, 1988). When we express both the BolA gene as well as silicatein, we are able to construct round cells, coated in glass.
Methods
The phenotype of the cells expressing BolA is very different from the phenotype of wildtype E. coli. If the gene is successfully overexpressed, the cells become round, which we can easily observe under a widefield microscope. In widefield microscopy, the whole sample is simultaneously illuminated using a white light source so the phenotype of the sample can be inspected. This is comparable to normal light microscopy.
In order to obtain round cells we tested transforming E. coli BL21 with BolA under both an inducible promoter (Lac) and a constitutive promoter (J23100). Furthermore, we tried if transforming a strain with both BolA and the OmpA-silicatein fusion plasmid yielded round, glass covered cells. The following strains and conditions were tested under the widefield microscope:
| Plasmid(s) | IPTG | Silicic acid |
|---|---|---|
| Lac-BolA (inducible) | - | - |
| Lac-BolA (inducible) | + | - |
| J23100-BolA (constitutive) | - | - |
| OmpA-Silicatein (T. aurantia) + Lac-BolA (inducible) | + | + |
The cells were heat-fixed on a slide and observed under the widefield microscope.
Results and discussion
The four different strains were imaged under the widefield microscope, the taken images are shown in figure 2.
In figure 2, the widefield images of the four tested strains are shown. We can see from figure 2A that solely transforming E. coli with BolA but not inducing the plasmid results in cells with the phenotype of wildtype E. coli; the cells are rod-shaped. Figure 2B shows that induction of the cells transformed wil BolA under the inducible Lac-promoter indeed has changed the phenotype of the cell. The cells have clearly become spherical. Constitutive expression of the BolA gene, as shown in 1C has the opposite result: the cells are rod-shaped and elongated. This is probably because of the stress the plasmid puts on the cells. As mentioned before, BolA is a gene involved in the stress response of E. coli that changes the morphology of the cells. A too high expression of the gene could therefore change the morphology in an unexpected way, such as elongation, a phenomena that is often observed in E. coli (Höltje, 1998). Since we require sphere-shaped cells, constitutive expression of the BolA-gene is not desired. Co-expression of the OmpA-silicatein fusion plasmid and the inducible BolA plasmid also yielded round cells, as seen in figure 2D.
So, by inducing the expression of BolA, we are indeed able to control the shape of E. coli and turn the cell into a sphere. Also, transforming a cell with both silicatein-OmpA and BolA yields round cells, so the formation of the glass layer does not distort the cell shape. The glass layer can not be seen under the widefield microscope, but we previously confirmed the presence of the silica layer with AFM and rhodamine staining. Being able to control cell shape is of major importance if we want to create biological lenses, since the lenses are desired in various sizes and shapes. Especially spherical lenses are useful since they are symmetrical and therefore do not require a specific orientation; they will focus the light in the same way whatever their orientation is. From figure 2 we can see that the ells are not perfectly homogeneously shaped, there is some variation between the shape of the different cells. This variation is even clearer for the cells that contain the silica layer. This is possibly because two plasmids with a lac promoter put a great strain on the cells, resulting in a greater variation. For precision optics, it is extremely important that there is little to none variation between the lenses. Therefore, it’s recommended to do more research in controlling cell shape. However, since there is always a variation in gene expression between cells it will be wiser to conduct research into selecting or sorting the cells on their size or shape. A possible solution could be sorting the cells using FACS. As a future experiment, we could stain the silica-coated cells with Rhodamine and let a FACS-machine sort the cells on their phenotype.
Back to TopSterilization of the biolenses
Place experiment (Introduction, methods, results&Discussion here)
Back to TopImproving solar cells with biolenses
Place experiment (Introduction, methods, results&Discussion here)
Back to TopDiscussion & Conclusions
Conclusion and discussion on the experiments
Conclusions & Recommendations
References
- Aldea, M., Hernandez-Chico, C., De La Campa, A., Kushner, S., & Vicente, M. (1988). Identification, cloning, and expression of bolA, an ftsZ-dependent morphogene of Escherichia coli. Journal of bacteriology, 170(11), 5169-5176.
- Cha, J. N., Shimizu, K., Zhou, Y., Christiansen, S. C., Chmelka, B. F., Stucky, G. D., & Morse, D. E. (1999). Silicatein filaments and subunits from a marine sponge direct the polymerization of silica and silicones in vitro. Proceedings of the National Academy of Sciences, 96(2), 361-365.
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