Difference between revisions of "Team:TU Delft/Header"

 
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        <title>iGEM TU Delft</title>
 
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    <body id="delft">
 
  
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        <link rel="stylesheet" href="https://2016.igem.org/Template:TU_Delft/Bootstrap?action=raw&ctype=text/css">
  
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         <div class="page-heading text-center">
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        <link rel="stylesheet" href="https://2016.igem.org/Template:TU_Delft/FA?action=raw&ctype=text/css">
  
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            <div class="container">
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        <link rel="stylesheet" href="https://2016.igem.org/Template:TU_Delft/Style?action=raw&ctype=text/css">
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      #top_title, #sideMenu{
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                <h1 class="page-header">Opticoli<span class="title-under"></span></h1>
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      #content{
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                <h3> The new age of optics: Producing biological lenses and lasers to improve microscopy </h3>
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    </head>
  
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        <div  class="main-container project">
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    <body>
  
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            <div class="container">
 
  
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    <header class="main-header">       
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                <div class="our-project">
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            <div class="navbar-top">  
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                    <span class="anchor" id="description"></span>
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                    <h2 class="title-style-1">Project Description<span class="title-under"></span></h2>
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                    <div class="col-sm-6 col-xs-12">
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                <div class="row">
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                    <div class="col-md-6 biolaser">
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                        <ul class="list-unstyled list-inline header-contact">
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                        <h2 class="title-style-2">BIOLASERS</h2>
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                            <li> <i class="fa fa-envelope"></i> <a href="mailto:tudelft.igem@gmail.com">tudelft.igem@gmail.com</a> </li>
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                        <p> <strong>Imaging cells </strong>is essential for understanding life at the smallest scale and fighting cellular diseases like cancer. Imaging often relies on <strong>fluorescence</strong>, but fluorescent proteins have some drawbacks, such as their wide spectrum and low intensity.</p>
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                      </ul> <!-- /.header-contact  -->
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                        <p> Our <strong>biolasers</strong> will provide an accurate, safe and biological way to improve this.</p>
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                        <p> 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.<strong> Lasing</strong> works with the same principle as fluorescence, but now the light source is put between <strong>mirrors</strong>. The photons keep <strong>‘bouncing’</strong>, increasing the energy of the system. When the light gets a certain power, the photons can escape in the form of a laser beam.</p>                           
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                        <div  class="project-col">
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                            <img src="https://static.igem.org/mediawiki/2016/8/8a/TU_Delft_Lasingfull.png" alt="laser">
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                        </div>
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                        <p>A biolaser is achieved by trapping fluorescent proteins inside a <strong>reflective agent</strong>. We have chosen two reflective agents: bioglass (polysilicate) and bioplastic (PHB). By covering a cell with <strong>polysilicate</strong>, the photons can resonate inside the cell, making a whole-cell laser. The polysilicate is synthesized by an enzyme called <strong>silicatein</strong>, which is expressed on the cell wall by fusion to membrane proteins. By filling a cell with <strong>PHB</strong>, 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 <strong>PHB granules</strong>.</p>                                     
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                        <div class="project-col col-md-6 col-sm-6">
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                            <img src="https://static.igem.org/mediawiki/2016/b/bd/TU_Delft_frontlaser.png" alt="whole cell laser">
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                        <div class="project-col col-md-6 col-sm-6">
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                                <img src="https://static.igem.org/mediawiki/2016/c/cc/TU_Delft_singlecelllaser.png" alt="intracellular laser">
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                            </div>
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                        <h2 class="title-style-2 biolens">BIOLENSES</h2>                       
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                        <p><strong>Microlenses</strong> 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 <strong>environment</strong>. </p>
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                        <p>Our biological microlens will be cheap, easy to make and environmentally friendly.</p>
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                        <p>When we cover a cell with <strong>polysilicate</strong>, using the enzyme <strong>silicatein</strong>, 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 <strong>morphology</strong> and investigate <strong>optical properties</strong>. These enlarged cells can also be used in the lasing experiments. The single cell will be able to diffract light as a <strong>single microlens</strong>. When we make a grid of lenses, a <strong>microlens array</strong>, we can use the lens for a coating for solar panels, thin lightweight cameras with high resolution or 3D screens.</p>
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                        <div class="project-col col-md-6 col-sm-6">
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                            <img src="https://static.igem.org/mediawiki/2016/7/71/TU_Delft_frontlens.png" alt="lenses">
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                        </div>
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                            <img src="https://static.igem.org/mediawiki/2016/3/37/TU_Delft_longcellroundcell.png" alt="long cell round cell">
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                <div class="our-project">
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                    <span class="anchor" id="results"></span>
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                    <h2 class="title-style-1">Experiments and results<span class="title-under"></span></h2>
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                    <!-- -Short introduction on our experiments and methods here --> 
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                    <span class="anchor" id="fluorophores"></span>
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                    <h2 class="title-style-2">Expression of different fluorophores</h2>
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                    <h3>  Introduction </h3> 
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                    <p>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 <a href="#references">(Cormack <i>et al.</i>, 1996; Nagai <i>et al.</i>, 2002; Shcherboo <i>et al.</i>, 2007; Rizzo <i>et al.</i>, 2004)</a>.</p>
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                    <p>Since mVenus, mKate and mCerulean did not exist in the iGEM registry yet, we
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                        constructed a brand new part for each of these, including strong constitutive
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                        promoter, RBS and terminators. GFP, on the other hand, was present in a whole
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                        range of biobricks. However, to our knowledge, there was no single biobrick
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                        available containing promoter, RBS and terminators. Hence, we constructed a
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                        new biobrick containing all of the above using the existing part <b><a href="http://parts.igem.org/Part:BBa_E0840" target="_blank">E0840</a></b>,
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                        consisting of RBS, coding sequence and terminators. By means of PCR we amplified
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                        this biobrick with primers designed to add a promoter while mainaining the
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                        biobrick prefix and suffix. Not only did we express GFP under the strong
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                        promoter <b><a href="http://parts.igem.org/Part:BBa_J23100" target="_blank">J23100</a></b>, but also under less strong promoters
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                        <b><a href="http://parts.igem.org/Part:BBa_J23113" target="_blank">J23113</a></b>, <b><a href="http://parts.igem.org/Part:BBa_J23117" target="_blank">J23117</a></b>,
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                        <b><a href="http://parts.igem.org/Part:BBa_J23105" target="_blank">J23105</a></b>, and <b><a href="http://parts.igem.org/Part:BBa_J23108" target="_blank">J23108</a></b>.
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                        This way we were able to see the influence of promoter strength on
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                        fluorescent output.</p>
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                    <h3>  Experiments & Results </h3>
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                    <p>In order to characterize them before further use, the emission and absorption spectra were measured. Additionally, their effect on cell growth was investigated.</p>
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                                    <h4 class="panel-title">
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                                        Emission spectra of different fluorophores
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                                    </h4>
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                            </a>
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                                    <h3>Introduction</h3>
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                                    <p>To assure the fluorophores were functional, the emission spectra were recorded at the given excitation wavelength.</p>
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                                    <h3>Methods</h3>
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                                    <p>Parts encoding the four different fluorophores, including promoter, RBS and terminators, were expressed in <i>E. coli</i> 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. </p>
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                                    <p>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.  </p>
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                                    <p>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.
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                                        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.</p>
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                                    <h3>Results and Discussion</h3>
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                                    <strong>Fluorophores expressed under strong constitutive promoter</strong>
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                                    <p>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.</p>
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                                    <figure>
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                                        <center><img src="https://static.igem.org/mediawiki/2016/4/43/T--TU_Delft--Spectra_fluorophores.png" alt="fluorophores">
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                                            <figcaption><b>Figure 1,</b> Emission spectra of the fluorophores GFP, mVenus, mCerulean, and mKate expressed under strong promoter J23100. Excitation wavelength was 488 nm, 510 nm, 433 nm, 558 nm, respectively.</figcaption></center>
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                                    </figure>
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                                    <strong>GFP expressed under  constitutive promoters of different strengths.</strong>
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                                    <p>Not only did we express GFP under the strong promoter J23100, but also under less strong promoters J23113, J23117, J23105, and J23108.</p>
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                                    <figure>
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                                        <center><img src="https://static.igem.org/mediawiki/2016/3/32/T--TU_Delft--GFP_Spectra.png" alt="fluorophores">
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                                            <figcaption><b>Figure 2,</b> Emission spectra of GFP expressed under control of promoters with different strengths. Excitation wavelength was 488 nm.</figcaption></center>
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                                    </figure>
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                                    <p>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.</p>
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                                    <figure>
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                                        <center><img src="https://static.igem.org/mediawiki/2016/4/46/T--TU_Delft--Spectra_GFP_individual.png" alt="fluorophores">
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                                            <figcaption><b>Figure 3,</b> Emission spectra of GFP expressed under control of promoters with different strengths, normalized by 1. Excitation wavelength was 488 nm.</figcaption></center>
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                                    </figure>
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                                    <a href="#fluorophores" class="btn btn-info" role="button" onclick="$('html,body').animate({scrollTop: $('#fluorophores').offset().top}, 'slow');" style="text-decoration:none; color:#f3f4f4; float:right;">Back to Top</a>                                   
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                                </div>
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                                    <h4 class="panel-title">
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                                        The effect of expression of fluorophores on growth
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                                    </h4>
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                                    <h3>Introduction</h3>
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                                    <p>
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                                        As consitutive expression can sometimes be hard on the cells, we investigated
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                                        the effect of the different consitutive promoters on cell growth. In a 24 hour
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                                        kinetic cycle alterating shaking at 37°C with fluorescence and optical density
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                                        measurements, we investigated whether this was the case.
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                                    </p>
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                                    <h3>Methods</h3>
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                                    <p>
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                                        An overnight culture in eM9 medium was inoculated in fresh eM9 to an OD of 0.1
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                                        in a 96 well plate. The emission at 522 nm was measured every 15 minutes.
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                                        Measurements were done in quadruplicate with pure eM9 as a blank.
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                                    </p>
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                                    <h3>Results and Discussion</h3>
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                                    <p>
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                                        Figure 1 shows the 24 hour measurement of optical density and fluorescence
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                                        intensity. The final OD is approximately equal for all different strains,
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                                        suggesting that the level of constitutive expression was not influencing the
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                                        growth. Furthermore, fluorescence intensity drops after the exponential growth
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                                        phase, suggesting that GFP is being broken down by proteases as a response to
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                                        nutrient limitation. After this event, growth continues at a slower pace, while
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                                        GFP activity keeps decreasing. All in all, constitutive expression of GFP does
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                                        not seem to have a detrimental effect on cell growth during exponential phase.
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                                    </p>
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                                    <figure>
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                                        <center><img src="https://static.igem.org/mediawiki/2016/7/76/T--TU_Delft--Kinetic_cycle_fluorophores.png" alt="fluorophores">
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                                            <figcaption><b>Figure 1.</b> Kinetic measurement of fluorescence intensity at
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                                                522 nm and optical density at 600 nm, while shaking at 37°C. Above
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                                                6·10<sup>4</sup> the intensity was too high to be measured.</figcaption></center>
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                                    </figure>
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                                    <a href="#fluorophores" class="btn btn-info" role="button" onclick="$('html,body').animate({scrollTop: $('#fluorophores').offset().top}, 'slow');" style="text-decoration:none; color:#f3f4f4; float:right;">Back to Top</a>                                   
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                    <h3> Discussion & Conclusions </h3>
 
−
 
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                    <p>We were able to succesfully construct and characterize two biobricks with brand new fluorophores
 
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                        for the iGEM registry: <b><a href="http://parts.igem.org/Part:BBa_K1890011" target="_blank">mVenus</a></b>
 
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                        and <b><a href="http://parts.igem.org/Part:BBa_K1890010" target="_blank">mCerulean</a></b>.
 
−
                        Also, we constructed <b><a href="http://parts.igem.org/Part:BBa_K1890020" target="_blank">five new composite parts</a></b>,
 
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                        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,
 
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                        which can be very usefull for future iGEM teams.</p><br>
 
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                    <span class="anchor" id="silicatein"></span>
 
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                    <h2 class="title-style-2">Coating the cell in polysilica using silicatein</h2>
 
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                    <h3>  Introduction </h3> 
 
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                    <p>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. <i>E. coli</i> 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 <a href="#references">(Müller et al., 2008; Müller et al. 2003)</a>. Therefore, we are transforming <i>E. coli</i> with silicatein-α. We test the use of two different silicateins, one originating from the marine sponge <i>Suberites domuncula</i> <a href="#references">(Müller, 2011)</a> and one originating from the marine sponge <i>Tethya aurantia</i> <a href="#references">(Cha et al., 1999)</a>. We express the enzyme in three different ways. First of all, we expressed the gene from <i>S. domuncula </i> and see if the enzyme is transported outside the cell as described by <a href="#references">Müller et al, 2008</a>. Furthermore, we express a fusion of silicatein from <i>T. aurantia</i> to the trans-membrane protein OmpA (outer membrane protein A) from <i>E. coli</i> to anchor the silicatein to the membrane <a href=#"references">(Curnow, Kisailus, & Morse, 2006; Francisco et al. 1992)</a>, which might make coating the cell specifically in polysilicate more efficient. We also express a fusion of silicatein from <i>S. domuncula</i> to the transmembrane Ice Nucleation Protein (INP) from  <i>Pseudomonas syringae</i>, a popular protein for membrane fusions <a href="#references">(Kim & Yoo, 1998)</a>. 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. </p>
 
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                    <figure><center>
 
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                            <img src="https://static.igem.org/mediawiki/2016/2/29/T--TU_Delft--Silicatein_ovv.png" alt="Silicatein">
 
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                            <figcaption><b>Figure 1,</b> (A) Silicatein is able to convert monosilicate to polysilicate, allowing the cell to cover itself in polysilicate. (B) We express silicatein in three ways: solely expressing silicatein and fusing it to the membrane proteins OmpA or INP. </figcaption></center>
 
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                        <h3>  Experiments & Results </h3>
 
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−
                        <p>To determine if we have successfully covered <i>E. coli</i> 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.</p>
 
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                                            Rhodamine staining of silica covered cells
 
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                                        <h3>Introduction</h3>
 
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                                        <p>After transforming <i>E. coli</i> 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  <a href="#references">(Li, Chu, & Lee, 1989)</a>. 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. </p>
 
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                                        <h3>Methods</h3>
 
−
                                        <p> The experiment was performed using <i>E. coli</i> BL21 with the following plasmids and conditions. All genes are under an inducible promoter. </p>
 
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                                        <table class="notebook table table-style-1">
 
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                                            <thead>
 
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                                            <th>Plasmid(s)</th>
 
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                                            <th>IPTG</th>
 
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                                            <th>Silicic acid</th>
 
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                                            <th>Rhodamine 123</th>
 
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                                            </thead>
 
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                                            <tbody>
 
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                                                <tr>
 
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                                                    <td> OmpA-Silicatein </td>
 
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                                                    <td>+</td>
 
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                                                    <td>+</td>
 
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                                                    <td>+</td>
 
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                                                </tr>
 
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                                                <tr>
 
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                                                    <td> Silicatein  </td>
 
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                                                    <td>+</td>
 
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                                                    <td>+</td>
 
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                                                    <td>+</td>
 
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                                                <tr>
 
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                                                    <td>INP-Silicatein </td>
 
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                                                    <td>+</td>
 
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                                                    <td>+</td>
 
−
                                                    <td>+</td>
 
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                                                </tr>
 
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                                                <tr>
 
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                                                    <td>OmpA-Silicatein (negative control) </td>
 
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                                                    <td>+</td>
 
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                                                    <td>-</td>
 
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                                                    <td>+</td>
 
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                                                </tr>
 
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                                                <tr>
 
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                                                    <td>OmpA-Silicatein </td>
 
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                                                    <td>+</td>
 
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                                                    <td>+</td>
 
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                                                    <td>-</td>
 
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                                            </tbody>
 
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                                        </table>
 
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                                        <p>The cells were stained with 0.1 vol% Rhodamine. And washed 5 times with PBS, prior to imaging <a href="#references">(Li et al., 1989; Müller et al., 2005)</a>. Both widefield- (light microscopy) and fluorescence microscopy were used to image the cells.</p>
 
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                                        <h3>Results and discussion</h3>
 
−
                                        <p>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).</p>
 
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                                        <figure>
 
−
                                            <img src="https://static.igem.org/mediawiki/2016/6/65/T--TU_Delft--Silicatein9.png" alt="Rhodamine staining" >
 
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                                            <figcaption><b>Figure 1,</b> widefield and fluorescence images of OmpA-silicatein with silicic acid and OmpA-silicatein without silicic acid (negative control) at maximum excitation energy. Of the widefield and fluorescence images an overlay was made to show the fraction of fluorescent cells. The negative control causes overexposure of the camera, therefore the fluorescent image only gives one uniform signal. </figcaption>
 
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                                        </figure>
 
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                                        <p>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. </p>
 
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                                        <figure>
 
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                                            <img src="https://static.igem.org/mediawiki/2016/f/f9/T--TU_Delft--Silicatein3.png" alt="Rhodamine staining" >
 
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                                            <figcaption><b>Figure 2,</b> widefield and fluorescence images of silicatein with silicic acid , INP-silicatein with silicic acid and OmpA-silicatein without silicic acid (negative control) at one-third of the maximum excitation energy. Of the widefield and fluorescence images an overlay was made to show the fraction of fluorescent cells. </figcaption>
 
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                                        </figure>
 
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                                        <p>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.</p>
 
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                                        <a href="#silicatein" class="btn btn-info" role="button" onclick="$('html,body').animate({scrollTop: $('#silicatein').offset().top}, 'slow');" style="text-decoration:none; color:#f3f4f4; float:right;">Back to Top</a>                                   
 
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                                            Imaging of silicatein-expressing cells using SEM
 
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                                        <p> <!-- Replace this--> Place experiment (Introduction, methods, results&Discussion here)</p>
 
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                                            Imaging of silicatein-expressing cells using TEM
 
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                                            Analysis of physical properties of silica covered cells using AFM
 
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                                        <p> <!-- Replace this--> Place experiment (Introduction, methods, results&Discussion here)</p>
 
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                                            Viability experiments of silica covered cells
 
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                                        <h3>Introduction</h3>
 
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                                        <p>Since the silicatein expressing cells are to cover themselves in polysilicate, their nutrient supply might be limited by diffusion,
 
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                                            which can eventually result in cell death. To investigate whether this is indeed the case,a growth study was performed.</p>
 
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                                        <h3>Methods</h3>
 
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                                        <p>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
 
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                                            supplemented with silicic acid as substrate for silicatein. During the following five hours samples were taken, of which a 10<sup>-6</sup> dilution was
 
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                                            plated on selective LB plates. Colony forming units (cfu) were counted the day after.</p>
 
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                                        <h3>Results and Discussion</h3>
 
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                                        <p>Cells expressing either silicatein from <i>S. domuncula</i> (Sil Sdom), silicatein from <i>S. domuncula</i> fused to INP (INP Sil Sdom) or silicatein from <i>T. aurantia</i> fused to OmpA (OmpA Sil Taur) were tested.
 
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                                            As a negative control, OmpA Sil Taur expressing cells without silicic acid were used. After one hour no colonies were observed
 
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                                            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.</p>
 
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                                        <figure>
 
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                                            <center><img src="https://static.igem.org/mediawiki/2016/8/89/T--TU_Delft--viability_silicateins.png" alt="Hardware setup">
 
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                                                <figcaption><b>Figure 1,</b> Number of colony forming units (cfu) during incubation with silicic acid.</figcaption></center>
 
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                        <h3> Discussion & Conclusions </h3>
 
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                        <p> <!-- Replace this--> Conclusion and discussion on the experiments</p>
 
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                    <span class="anchor" id="Biolaser"></span>
 
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                    <h2 class="title-style-2">Engineering a biological laser</h2>
 
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                    <h3>  Introduction </h3> 
 
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                    <p>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 <a href="#references">(Gather & Yun, 2011; Svelto & Hanna, 1976)</a>. More information on this can be found in the <b><a href="#description">project description</a></b>. 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 <a href="#references">(Eggeling, Widengren, Rigler, & Seidel, 1998)</a>. 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 <a href="#references">(Jonáš et al., 2014)</a>. 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. </p>
 
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                    <figure>
 
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                        <center><img src="https://static.igem.org/mediawiki/2016/1/1f/T--TU_Delft--Setupfinal.jpg" alt="Hardware setup">
 
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                            <figcaption>Our custom-built hardware setup to image our Biolaser cells</figcaption></center>
 
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                    </figure>
 
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                    <p>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 <b><a href="https://2016.igem.org/Team:TU_Delft/Hardware" target="_blank">hardware page</a></b>.</p>
 
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                    <p> Using our custom-built setup, we analysed our ‘Biolaser’-cells, to see if they were able to produce a laser-like emission of light. </p>
 
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                    <h3>  Experiments & Results </h3>
 
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                    <p>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.</p>
 
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                                        Imaging our biolaser using our home-built setup
 
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                                    <h3>Introduction</h3>
 
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                                    <p>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. </p>
 
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                                    <figure><center>
 
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                                            <img src ="https://static.igem.org/mediawiki/2016/c/c0/T--TU_Delft--hardware2.png" alt = "setup">
 
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                                            <figcaption><b>Figure 1,</b> the design of our custom self-built setup</figcaption>
 
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                                        </center></figure>
 
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                                    <p>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.</p>
 
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                                    <h3>Methods</h3>
 
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                                    <p>In order to confirm whether the setup was working, we used <i> E. coli</i> 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.</p>
 
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                                    <p>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.</p>
 
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                                    <h3>Results & discussion</h3>
 
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                                    <p> Imaging the cells with the setup yielded the following results: </p>
 
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                                    <figure>
 
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                                        <img src="https://static.igem.org/mediawiki/2016/2/2a/T--TU_Delft--Cellsinsetup.png" alt="Setup results"  >
 
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                                        <figcaption> <b>Figure 2,</b> <i> E. coli</i> cells transformed with (A)OmpA-silicatein fusion plasmid and (B) mCerulean, imaged in our custom self-built optical setup. The cells were excited with a laser at a wavelength of 405nm and an intensity of 0.5 mW. </figcaption>
 
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                                    </figure>
 
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                                    <p>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. </p>
 
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                                    <p>For this experiment we used <i>E. coli</i> 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.</p>
 
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                                        Intensity measurements of laser cells
 
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                                    <h3>Introduction</h3>
 
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                                    <p>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.</p>
 
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                                    <center><figure>
 
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                                            <img src="https://static.igem.org/mediawiki/2016/6/6a/T--TU_Delft--Fluorescencevslasing.png" alt="Fluorescence vs. lasing">
 
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                                            <figcaption><b>Figure 1,</b> the relation of input energy vs. output intensity for fluorescence and lasing <a href="#references">(Fan & Yun, 2014)</a></figcaption>
 
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                                        </figure></center>
 
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                                    <p>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 </p>
 
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                                    <h3>Methods</h3>
 
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                                    <p>For this experiment we used <i>E. coli</i> BL21 transformed with the following BioBricks:</p>
 
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                                        <thead>
 
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                                        <th>Plasmid(s) and conditions</th>
 
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                                        <th>Function</th>
 
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                                        <tbody>
 
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                                            <tr>
 
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                                                <td> mCerulean (constitutively expressed) </td>
 
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                                                <td>Fluorescence</td>
 
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                                            <tr>
 
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                                                <td> mCerulean (constitutive) + OmpA-silicatein (induced, incubated in silicic acid)</td>
 
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                                                <td>Biolaser</td>
 
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                                            <tr>
 
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                                                <td> OmpA-silicatein (induced, incubated in silicic acid)</td>
 
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                                                <td>Negative control (no fluorescence)</td>
 
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                                            </tr>
 
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                                    </table>
 
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                                    <p>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 <a href="#references">(McCloy et al., 2014)</a>. </p>
 
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                                    <h3>Results and discussion</h3>
 
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                                    <p>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. </p>
 
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                                    <figure>
 
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                                        <img src="https://static.igem.org/mediawiki/2016/9/90/T--TU_Delft--lasersetupresults.png" alt="Setupresults" >
 
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                                        <figcaption><b>Figure 2,</b> intensity measurements of cells transformed with mCerulean or mCerulean and OmpA-silicatein. The measured emission intensity was plotted against the excitation power (black dots) along a prediction of the increase of intensity (blue line). Left of the intensity graphs, a picture of the imaged spot is shown.</figcaption>
 
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                                    </figure>
 
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                                    <p>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). </p>
 
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                                    <p>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 <a href="#references">(Fan & Yun, 2014)</a>. 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.</p>
 
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                                    <p>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. <strong><a href= https://2016.igem.org/Team:TU_Delft/Model >Modeling</strong></a> showed, that in a cavity as big as <i>E. coli</i> (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.</p>
 
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                    <h3> Discussion & Conclusions </h3>
 
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                    <p> 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.</p>
 
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                    <span class="anchor" id="Biolenses"></span>
 
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                    <h2 class="title-style-2">Engineering biological lenses</h2>
 
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                    <h3>  Introduction </h3> 
 
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                    <p> <!-- Replace this--> Introduction on lenses & experiments</p>
 
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                    <p> <!-- Replace this--> Introduction on the experiments that we did</p>
 
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                                        Influencing cell shape for round lenses
 
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                                    <h3>Introduction & background</h3>
 
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                                    <p>When making biological lenses, the shape of the lens is of crucial importance. <i>E. coli</i> is a rod-shaped organism, so it’s not symmetrical along all axes. Shining light on the round parts of <i>E. coli</i> 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 <b><a href="https://2016.igem.org/Team:TU_Delft/Model" target="_blank">modeling page</a></b>.</p>
 
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                                    <figure>
 
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                                        <center><img src="https://static.igem.org/mediawiki/2016/9/9c/T--TU_Delft--Breakinglight.png" alt="Diffraction">
 
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                                            <figcaption> <b>Figure 1:</b> Different ways of focusing light by rod-shaped lenses and spherical lenses. The rod-shaped lens has various directions in which it can break the light, therefore the orientation of this lens is extremely important. The spherical cell only has one, so the orientation of the cell does not matter.</figcaption></center>
 
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                                    </figure>
 
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                                    <p> 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 <i>E. coli</i> 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.</p>
 
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                                    <p>In order to create spherical E. coli, we overexpress the <i>BolA</i> gene. <i>BolA</i> is a gene that controls the morphology of <i>E. coli</i> in the stress response <a href="#references">(Santos, Freire, Vicente, & Arraiano, 1999)</a>. By overexpressing this gene, the rod-shaped <i>E. coli</i> cells will become round <a href="#references">(Aldea, Hernandez-Chico, De La Campa, Kushner, & Vicente, 1988)</a>. When we express both the <i>BolA</i> gene as well as silicatein, we are able to construct round cells, coated in glass.</p>
 
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                                    <h3>Methods</h3>
 
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                                    <p>The phenotype of the cells expressing BolA is  very different from the phenotype of wildtype <i>E. coli</i>. 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. </p>
 
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                                    <p> In order to obtain round cells we tested transforming <i>E. coli</i> BL21 with <i>BolA</i> under both an inducible promoter (Lac) and a constitutive promoter (J23100). Furthermore, we tried if transforming a strain with both <i>BolA</i> and the OmpA-silicatein fusion plasmid yielded round, glass covered cells. The following strains and conditions were tested under the widefield microscope:</p>
 
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                                        <thead>
 
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                                        <th>Plasmid(s)</th>
 
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                                        <th>IPTG</th>
 
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                                        <th>Silicic acid</th>
 
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                                        <tbody>
 
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                                                <td> Lac-<i>BolA</i> (inducible)  </td>
 
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                                                <td>-</td>
 
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                                                <td>-</td>
 
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                                            <tr>
 
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                                                <td> Lac-<i>BolA</i> (inducible)  </td>
 
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                                                <td>-</td>
 
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                                            <tr>
 
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                                                <td>J23100-<i>BolA</i> (constitutive)  </td>
 
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                                                <td>-</td>
 
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                                                <td>-</td>
 
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                                                <td>OmpA-Silicatein (<i>T. aurantia</i>) + Lac-<i>BolA</i> (inducible)  </td>
 
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                                                <td>+</td>
 
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                                                <td>+</td>
 
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                                    <p>The four different strains were imaged under the widefield microscope, the taken images are shown in figure 2.</p>
 
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                                            <img src="https://static.igem.org/mediawiki/2016/e/e6/T--TU_Delft--BolA_widefield.png" alt="BolA">
 
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                                            <figcaption><b> Figure 2: </b> Widefield images of <i>E.coli</i> BL21 transformed with <b>(A)</b> <i>BolA</i> under the inducible Lac-promoter, uninduced, <b>(B)</b> <i>BolA</i> under the inducible Lac promoter and induced with 1 mM IPTG, <b>(C)</b> <i>BolA</i> under the constitutive promoter J23100, <b>(D)</b> <i>BolA</i> and OmpA-silicatein fusion under the inducible Lac promoter and induced with 1 mM IPTG.  </figcaption></center>
 
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                                    </figure>
 
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                                    <p> In figure 2, the widefield images of the four tested strains are shown. We can see from figure 2A that solely transforming <i>E. coli</i> with <i>BolA</i> but not inducing the plasmid results in cells with the phenotype of wildtype <i>E. coli</i>; the cells are rod-shaped. Figure 2B shows that induction of the cells transformed wil <i>BolA</i> under the inducible Lac-promoter indeed has changed the phenotype of the cell. The cells have clearly become spherical. Constitutive expression of the <i>BolA</i> 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,  <i>BolA</i> is a gene involved in the stress response of <i>E. coli</i> 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 <i>E. coli</i> (Höltje, 1998). Since we require sphere-shaped cells, constitutive expression of the <i>BolA</i>-gene is not desired. Co-expression of the OmpA-silicatein fusion plasmid and the inducible <i>BolA</i> plasmid also yielded round cells, as seen in figure 2D.</p>
 
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                                    <p>So, by inducing the expression of <i>BolA</i>, we are indeed able to control the shape of <i>E. coli</i> and turn the cell into a sphere. Also, transforming a cell with both silicatein-OmpA and <i>BolA</i> 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. </p>
 
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                    <h2 class="title-style-1">Conclusions & Recommendations<span class="title-under"></span></h2>
 
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            <h4 class="footer-title">References</h4>
 
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                <li>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. </li>
 
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                <li>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. </li>
 
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                <li>Cormack, B. P., Valdivia, R. H., & Falkow, S. (1996). FACS-optimized mutants of the green fluorescent protein (GFP). Gene, 173(1), 33-38.</li>
 
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                <li>Curnow, P., Bessette, P. H., Kisailus, D., Murr, M. M., Daugherty, P. S., & Morse, D. E. (2005). Enzymatic synthesis of layered titanium phosphates at low temperature and neutral pH by cell-surface display of silicatein-?? Journal of the American Chemical Society, 127(45), 15749–15755.</li>
 
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                <li>Curnow, P., Kisailus, D., & Morse, D. E. (2006). Biocatalytic Synthesis of Poly (L‐Lactide) by Native and Recombinant Forms of the Silicatein Enzymes. Angewandte Chemie International Edition, 45(4), 613-616. </li>
 
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                <li>Eggeling, C., Widengren, J., Rigler, R., & Seidel, C. (1998). Photobleaching of fluorescent dyes under conditions used for single-molecule detection: Evidence of two-step photolysis. Analytical Chemistry, 70(13), 2651-2659. </li>
 
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                <li>Fan, X., & Yun, S.-H. (2014). The potential of optofluidic biolasers. Nature methods, 11(2), 141-147. </li>
 
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                <li>Francisco, J. a, Earhart, C. F., & Georgiou, G. (1992). Transport and anchoring of beta-lactamase to the external surface of Escherichia coli. Proceedings of the National Academy of Sciences of the United States of America, 89(April), 2713–2717.</li>
 
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                <li>Gather, M. C., & Yun, S. H. (2011). Single-cell biological lasers. Nature Photonics, 5(7), 406-410. </li>                   
 
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                <li>Höltje, J.-V. (1998). Growth of the stress-bearing and shape-maintaining murein sacculus of Escherichia coli. Microbiology and Molecular Biology Reviews, 62(1), 181-203. </li>
 
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                <li>Jonáš, A., Aas, M., Karadag, Y., Manioğlu, S., Anand, S., McGloin, D., Kiraz, A. (2014). In vitro and in vivo biolasing of fluorescent proteins suspended in liquid microdroplet cavities. Lab on a Chip, 14(16), 3093-3100. </li>
 
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                <li>Kim, E.-J., & Yoo, S.-K. (1998). Cell surface display of CD8 ecto domain on Escherichia coli using ice nucleation protein. Biotechnology techniques, 12(3), 197-201.</li>
 
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                <li>Li, C.-W., Chu, S., & Lee, M. (1989). Characterizing the silica deposition vesicle of diatoms. Protoplasma, 151(2-3), 158-163. </li>
 
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                <li>McCloy, R. A., Rogers, S., Caldon, C. E., Lorca, T., Castro, A., & Burgess, A. (2014). Partial inhibition of Cdk1 in G2 phase overrides the SAC and decouples mitotic events. Cell Cycle, 13(9), 1400-1412. </li>
 
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                <li>Müller, W. E. G. (2003). Silicon biomineralization.</li>
 
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                <li>Müller, W. E. G., Rothenberger, M., Boreiko, A., Tremel, W., Reiber, A., & Schröder, H. C. (2005). Formation of siliceous spicules in the marine demosponge Suberites domuncula. Cell and Tissue Research, 321(2), 285–297.</li>
 
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                <li>Müller, W. E., Engel, S., Wang, X., Wolf, S. E., Tremel, W., Thakur, N. L., Schröder, H. C. (2008). Bioencapsulation of living bacteria (Escherichia coli) with poly (silicate) after transformation with silicatein-α gene. Biomaterials, 29(7), 771-779. </li>
 
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                <li>Müller, W. E. (2011). Molecular biomineralization: aquatic organisms forming extraordinary materials: Springer Science & Business Media.</li>
 
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                <li>Nagai, T. et al. A variant of yellow fluorescent protein with fast and efficient maturation for cell-biological applications. Nat. Biotechnol. 20, 87–90 (2002).</li>
 
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                <li>Rizzo, M. A., Springer, G. H., Granada, B. & Piston, D. W. An improved cyan fluorescent protein variant useful for FRET. Nat. Biotechnol. 22, 445–449 (2004).</li>
 
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                <li>Santos, J. M., Freire, P., Vicente, M., & Arraiano, C. M. (1999). The stationary‐phase morphogene bolA from Escherichia coli is induced by stress during early stages of growth. Molecular microbiology, 32(4), 789-798. </li>
 
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                <li>Shcherbo, D., Merzlyak, E. M., Chepurnykh, T. V, Fradkov, A. F., Ermakova, G. V, Solovieva, E. a, … Chudakov, D. M. (2007). Bright far-red fluorescent protein for whole-body imaging. Nature Methods, 4(9), 741–746. http://doi.org/10.1038/nmeth1083</li>
 
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                <li>Svelto, O., & Hanna, D. C. (1976). Principles of lasers: Springer.</li>
 
  
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Latest revision as of 18:25, 21 November 2016