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 promoter

As 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.

GFP expressed under constitutive promoters of different strengths.

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.

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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.

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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.

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Introduction

Electron microscopy is an imaging technique that can acquire images with resolutions far above the optical limit due to the small wavelength of electrons (Nellist, 2011). An electron gun forms an electron beam which is focused and accelerated by magnetic lenses. When the electrons reach the sample, there are several interactions possible with the sample (figure 1). These various interactions can be used for different techniques. We have used high-angle annular dark-filed transmission electron microscopy (HAAFD-TEM) to make an image of the sample, energy dispersive x-ray spectroscopy to analyze the element composition of the sample and scanning electron microscopy (SEM) to analyze the phenotype of the samples.

In HAAFD-TEM electrons pass through the sample and are inelastically scattered at a high angle and acquired by a detector. The amount of electrons scattered at a high angle depends on the thickness and the material of the sample. When the thickness is high, the sample scatters a lot it will appear bright in the image.

Energy dispersive x-ray spectroscopy (EDX) is an analytical technique to determine the elements present in the sample (Friel et al. ,2006). When the electron beam interacts with the sample x-rays can emerge from the sample. These electrons can be detected and are characteristic for the element which emitted the x-ray. In that way we can specifically determine which elements are present in our sample.

Methods

The experiment were performed using E.coli BL21 cells with the plasmid containing OmpA-Silicatein. Two samples were made where the first sample was induced with IPTG but no silicic acid was added and a second sample which was both induced with IPTG and silicic acid was added, therefore this sample will have a polysilicate layer. The samples were prepared by fixation using 1% polylysine on a quantifoil carbon grid.

Results and discussion

Both samples were imaged using HAAFD-TEM and energy dispersive x-ray spectroscopy (figure 2). The cells are fixed at a carbon grid. In each sample, we measured the silicon content. We can clearly see that the grid contains silicon and there is a lot of silicon signal from the grid (figure 2 B,D), since each blue dot represents silicon being measured. When a cell is positioned at a hole on in the grid we there is no background silicon signal, so we can observe the elemental composition of our sample. For the sample where no silicic acid is added, we can see some silicon present at the cell position. However, there is a significant increase in silicon detected for the sample where silicic acid was added to the sample. This shows that silicon co-localizes with the cell which means there is indeed a polysilica layer formed by the bacteria.

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Introduction

Methods

The polysilicate layer around the cells was prepared according to the polysilicate layer protocol. As a negative control, uninduces cultures (no IPTG) and cultures without substrate (no sodium silicate) were used. The samples were fixed with gluteraldehyde and imaged with SEM.

Results and Discussion

Figure 1 shows SEM images taken of samples in the presence (A, B) or absence (C, D) of silic acid. First of all, since we know from the rhodamine staining experiment that the polysilicate layer is present, it does not seem to influence the cell shape. However, the cells that are expected to have a polysilicate layer appear to be somewhat fused together. According to Müller et al (2008), cells possessing a polysilicate layer appear to be fused by a viscous cover. This can, however, also be the result of limited imaging resolution. When using titanium oxide as a substrate, Curnow et al (2005) reported large aggregates visible by SEM. This was not observed in the current experiment. Therefore, we can conclude that the polysilicate layer does not form aggregated or influence the shape of the cell, but form a homogeneous layer around the cell.

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Introduction

Due to encapsulation of the cell with a layer of polysilica the stiffness could change. We wanted to know whether and how the stiffness changes once e call is coated in polysilicate and determined this using Atomic Force Microscopy (AFM).

AFM is a technique used to determine surface characteristics of a sample. In AFM a very sharp tip scans over the surface of the sample. While scanning over the surface every height change is detected by a laser which is reflected by the cantilever on a position-sensitive photodetector (Figure 1). In the PeakForce QNM mode, ehich we used, also surface characteristics like the stiffness and deformation can be determined. The stiffness is computed from the relation between the force at which the cantilever pushes on the sample, the adhesion force and the deformation of the material.

Methods

Experiments were performed using E.coli BL21 cells transformed with the plasmid containing the OmpA-Silicatein gene. Silicic acid was added to one sample to form the polysilicate layer around the cell. Another sample with no silicic acid added was used as a control. The cells weres spun down and resuspended in MilliQ and fixated on a glass slide using 1% ABTES.

Results and discussion

We have imaged two samples with AFM. The first sample is OmpA fused with silicatein with silic acid added so that the cell can encapsulate itself with biosilica (figure 2A-B). The second sample, that wa used as a control, did not contain silicic acid (figure 2C-D). From both samples a height map (figure 2A,C) and the stiffness (figure 2B,D) was determined. Both samples were fixed on a glass slide in the same way. Due to a tip change is there a factor 10 difference between the stiffness of the glass slide of both samples.

Multiple cells were imaged and the relative stiffness of the cell compared to the stiffness of the glass slide was determined (figure 3). We found that cells covered with a layer of polysilicate have a stiffness of 0.43 compared to the glass slide and the cells without a layer of polysilicate have a stiffness of 0.16 compared to the glass slide. We can see that due to the encapsulation of the cells in polysilica the stiffness of the cells increases significantly.

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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.

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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.

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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.

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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). We will express this gene both under a constitutive promoter (Part K1890031), as well as an inducible promoter (Part K1890030). 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.

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Introduction

We wanted to test our biolenses in a real-world application: solar cells. This was chosen, since it was shown to make solar cells more efficient while using microlenses (Jutteau et al., 2015). To do these experiments, Dr. Arno Smets, professor Photovoltaic Materials and Devices of the TU Delft had sent us to Stefaan Heirman, a technician at the same research group, who helped us. However, our biolenses should leave the ML-1 lab, as Stefaan was located at another faculty in Delft. If in the future the biolenses will be used on solar panels, they are in the environment as well. Since we want to be able to implement our biolenses in a real-world application, the biolenses needed to be sterilized.

To sterilize, different options are available. For example alcohols, chlorine and chlorine compounds are used within healthcare and autoclaving is a much-used method in laboratory settings. Besides that, UV sterilization is a much-used method in water treatment (Chang et al., 1985; Hijnen et al., 2006; O’Flaherty et al., 2016). This is used to remove among others E. coli. Since we do not want any chemicals to potentially interfere with our biolenses, we have chosen not to use any of these chemicals and to test whether autoclaving or UV sterilization would be a good option.

Methods

Autoclaving biolenses

Aliquots of cells were autoclaved before imaged with scanning electron microscopy (SEM).

UV sterilization

A volume of 30 µL E. coli BL21 cells that were transformed with BolA in pBbA5c was plated on LB plates complemented with chloramphenicol. As controls for sterilization aliquots of 500 µL of the same cells were either heated at 95 °C for 10 minutes or washed with 500 µL 70% ethanol for three times. After sterilization, also 30 µL of these cells were plated on LB+Cm plates. Besides that, 30 µL of non sterilized cells were plated as well.

Two of the plates containing BolA transformed cells were UV sterilized in a Stratagene UV Stratalinker 1800. The manufacturer recommended power settings of 120000 µJ. In our experiment we have used a power of 120000 µJ and 240000µJ to sterilize our cells.

All plates were incubated overnight at 37 °C.

Results

Autoclaving

Cells containing the Sil_Sdom gene were autoclaved and subsequently imaged with SEM.

Since the SEM showed the cells were collapsed, and in no way could function as biolenses, another sterilization method was used.

UV sterilization

To test the effectivity of UV sterilization on our E. coli cells, a growth study after UV sterilization was performed. For this 30 µL BL21 cells transformed with BolA in pBbA5c was plated on LB plates complemented with chloramphenicol. These plates were UV sterilized in a Strategene UV Stratalinker 1800. As positive controls the same cells were heated for 10 minutes at 95 ºC or washed three times with 500 µL ethanol. As negative control the same cells were plated on a selective plate.

None of the sterilized plates showed any growth, while the non sterilized plate did yield colonies.

Besides the growth study, the cells were also imaged with SEM, which can be found in figure 2.

As shown in figure 2, the with silicate capturing of the cells remain undamaged after UV sterilization and could be used as biolenses now.

Discussion

In this experiment, different methods to sterilize our biolenses were tested, as we wanted to be able to put our biological microlenses in real-world conditions. For this, we have both autoclaved and UV sterilized our cells. From the growth study, it can be concluded that the biolenses were successfully sterilized by UV sterilization. Besides that, the shape of the biolenses was not affected by UV sterilization as opposed to autoclaving.

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Introduction

Ways to measure the efficiency of photovoltaics

There are different ways to measure the output of photovoltaic (PV) cells with accuracy and repeatability. The most used characteristic of PVs is their efficiency (Power Conversion Efficiency or PCE). It is defined as the energy produced from the device by the intended radiation from a light source. In order to measure the efficiency and produce acceptable results; a set protocols were set in the scientific community.

Those protocols are relevant for the intended radiation and the measurements taken. For the intended radiation we make the assumptions of air mass 1.5 (dictates how the sun’s radiation is influenced by the atmosphere), the measured surface is 37 degrees from the equator and 48.2 degrees from Zenith. Finally, the radiation is 1000Wm^-2 (Snaith, 2012) .

The parameters that influence the PCE measurements can be found in characteristic J-V diagrams, plotting current density (A/cm2) against voltage (V) (figure 1). The graph contains the three basic parameters of PCE measurements. Those are the short circuit current (I_sc), the open circuit voltage (V_oc) and the Fill Factor (FF). This graph is obtained by scanning the photovoltaic’s current output within a range of voltages, usually from -1 Volts to +1 Volts. The V_oc is the voltage when the current is zero and the I_sc is the current when the voltage is zero. The Fill Factor represents how much power the solar cell is producing or its theoretical maximum. The theoretical maximum power for each solar cell is \( P_{ideal} = V_{oc} \times I_{sc} \) and the maximum power is \( P_{max} = V_{max} \times I_{max}\) then the Fill Factor is defined as \(FF = \frac{P_{max}}{P_{ideal}} \). The Fill Factor is a defining term of the behavior of the solar cells (Grossiord, Kroon, Andriessen, & Blom, 2012).

Equipment and proposed measurements

In order to achieve those reproducible and accurate measurement of efficiency a certain type of equipment complemented with previously mentioned protocols need to be used. The equipment used to simulate the aforementioned conditions is called Solar Simulator, which is basically a big Xenon lamp with well-defined light power output. In order to calibrate the simulator a ‘standard’ cell is used with well-defined properties to measure the exact output of the simulator in each measurement. Finally, the tested device must be placed perpendicular to the light with nothing restricting the light path and not moved during the whole testing. Using this set of equipment, we are sure that we conduct all the experiments under the same scientifically accepted conditions so we remove a factor of uncertainty to our experimental results.

Methods

Results

Discussion

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