We would like to thank our student researchers over the years. In particular, Nina Meyer, Sibylle Mayer, Juliane Kaim, and Laura Jaggy, whose data have been utilized in this protocol as representative results. Research of our group presented here was funded by grants from the Deutsche Forschungsgemeinschaft (Hu 839/2-4, Hu 839/7-1) to Armin Huber.

<ol>
	<li>Wang, X., Huang, T., Bu, G., Xu, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dysregulation+of+protein+trafficking+in+neurodegeneration.">Dysregulation of protein trafficking in neurodegeneration.</a> Molecular Neurodegeneration. 9. 9, 31 (2014).</li><li>Schopf, K., Huber, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Membrane+protein+trafficking+in+Drosophila+photoreceptor+cells.">Membrane protein trafficking in Drosophila photoreceptor cells.</a> European Journal of Cell Biology. 96 (5), 391-401 (2017).</li><li>Hardie, R. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+photoreceptor+array+of+the+dipteran+retina.">The photoreceptor array of the dipteran retina.</a> Trends in Neurosciences. 9, 419-423 (1986).</li><li>Wang, T., Montell, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phototransduction+and+retinal+degeneration+in+Drosophila.">Phototransduction and retinal degeneration in Drosophila.</a> Pfl&#252;gers Archiv - European Journal of Physiology. 454 (5), 821-847 (2007).</li><li>Xiong, B., Bellen, H. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rhodopsin+homeostasis+and+retinal+degeneration:+lessons+from+the+fly.">Rhodopsin homeostasis and retinal degeneration: lessons from the fly.</a> Trends in Neurosciences. 36 (11), 652-660 (2013).</li><li>B&#228;hner, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Light-regulated+subcellular+translocation+of+Drosophila+TRPL+channels+induces+long-term+adaptation+and+modifies+the+light-induced+current.">Light-regulated subcellular translocation of Drosophila TRPL channels induces long-term adaptation and modifies the light-induced current.</a> Neuron. 34 (1), 83-93 (2002).</li><li>Cronin, M. A., Lieu, M. -. H., Tsunoda, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two+stages+of+light-dependent+TRPL-channel+translocation+in+Drosophila+photoreceptors.">Two stages of light-dependent TRPL-channel translocation in Drosophila photoreceptors.</a> Journal of Cell Science. 119, 2935-2944 (2006).</li><li>Meyer, N., Joel-Almagor, T., Frechter, S., Minke, B., Huber, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Subcellular+translocation+of+the+eGFP-tagged+TRPL+channel+in+Drosophila+photoreceptors+requires+activation+of+the+phototransduction+cascade.">Subcellular translocation of the eGFP-tagged TRPL channel in Drosophila photoreceptors requires activation of the phototransduction cascade.</a> Journal of Cell Science. 119, 2592-2603 (2006).</li><li>Oberegelsbacher, C., Schneidler, C., Voolstra, O., Cerny, A., Huber, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Drosophila+TRPL+ion+channel+shares+a+Rab-dependent+translocation+pathway+with+rhodopsin.">The Drosophila TRPL ion channel shares a Rab-dependent translocation pathway with rhodopsin.</a> European Journal of Cell Biology. 90 (8), 620-630 (2011).</li><li>Wagner, K., Smylla, T. K., Lampe, M., Krieg, J., Huber, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phospholipase+D+and+retromer+promote+recycling+of+TRPL+ion+channel+via+the+endoplasmic+reticulum.">Phospholipase D and retromer promote recycling of TRPL ion channel via the endoplasmic reticulum.</a> Traffic. , (2021).</li><li>Franceschini, N., Kirschfeld, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Les+ph&#233;nom&#233;nes+de+pseudopupille+dans+l'oeil+compose+de+Drosophila.">Les ph&#233;nom&#233;nes de pseudopupille dans l'oeil compose de Drosophila.</a> Kybernetik. 9 (5), 159-182 (1971).</li><li>Pichaud, F., Desplan, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+visualization+approach+for+identifying+mutations+that+affect+differentiation+and+organization+of+the+Drosophila+ommatidia.">A new visualization approach for identifying mutations that affect differentiation and organization of the Drosophila ommatidia.</a> Development. 128 (6), 815-826 (2001).</li><li>Smylla, T. K., Wagner, K., Huber, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Application+of+fluorescent+proteins+for+functional+dissection+of+the+Drosophila+visual+system.">Application of fluorescent proteins for functional dissection of the Drosophila visual system.</a> International Journal of Molecular Sciences. 22 (16), (2021).</li><li>Meyer, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanisms+of+the+light-dependent+translocation+of+the+ion+channel+TRPL+in+the+photoreceptors+of+Drosophila+melanogaster:+Dissertation+for+obtaining+the+academic+degree+Dr.+rer.+nat.">Mechanisms of the light-dependent translocation of the ion channel TRPL in the photoreceptors of Drosophila melanogaster: Dissertation for obtaining the academic degree Dr. rer. nat.</a> University of Karlsruhe (TH). , (2005).</li><li>Meyer, N., Oberegelsbacher, C., D&#252;rr, T. D., Sch&#228;fer, A., Huber, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+eGFP-based+genetic+screen+for+defects+in+light-triggered+subcelluar+translocation+of+the+Drosophila+photoreceptor+channel+TRPL.">An eGFP-based genetic screen for defects in light-triggered subcelluar translocation of the Drosophila photoreceptor channel TRPL.</a> Fly. 2 (1), 36-46 (2008).</li><li>Zheng, L., Carthew, R. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lola+regulates+cell+fate+by+antagonizing+Notch+induction+in+the+Drosophila+eye.">Lola regulates cell fate by antagonizing Notch induction in the Drosophila eye.</a> Mechanisms of Development. 125 (1-2), 18-29 (2008).</li><li>Hibbard, K. L., O'Tousa, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+role+for+the+cytoplasmic+DEAD+box+helicase+Dbp21E2+in+rhodopsin+maturation+and+photoreceptor+viability.">A role for the cytoplasmic DEAD box helicase Dbp21E2 in rhodopsin maturation and photoreceptor viability.</a> Journal of Neurogenetics. 26 (2), 177-188 (2012).</li><li>Huang, Y., Xie, J., Wang, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Fluorescence-based+genetic+screen+to+study+retinal+degeneration+in+Drosophila.">A Fluorescence-based genetic screen to study retinal degeneration in Drosophila.</a> PloS One. 10 (12), 0144925 (2015).</li><li>Zhao, H., Wang, J., Wang, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+V-ATPase+V1+subunit+A1+is+required+for+rhodopsin+anterograde+trafficking+in+Drosophila.">The V-ATPase V1 subunit A1 is required for rhodopsin anterograde trafficking in Drosophila.</a> Molecular Biology of the Cell. 29 (13), 1640-1651 (2018).</li><li>Xiong, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ER+complex+proteins+are+required+for+rhodopsin+biosynthesis+and+photoreceptor+survival+in+Drosophila+and+mice.">ER complex proteins are required for rhodopsin biosynthesis and photoreceptor survival in Drosophila and mice.</a> Cell Death and Differentiation. 27 (2), 646-661 (2020).</li><li>Zhao, H., Wang, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=PE+homeostasis+rebalanced+through+mitochondria-ER+lipid+exchange+prevents+retinal+degeneration+in+Drosophila.">PE homeostasis rebalanced through mitochondria-ER lipid exchange prevents retinal degeneration in Drosophila.</a> PLoS Genetics. 16 (10), 1009070 (2020).</li><li>Cerny, A. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+GTP-+and+phospholipid-binding+protein+TTD14+regulates+trafficking+of+the+TRPL+ion+channel+in+Drosophila+photoreceptor+cells.">The GTP- and phospholipid-binding protein TTD14 regulates trafficking of the TRPL ion channel in Drosophila photoreceptor cells.</a> PLoS Genetics. 11 (10), 1005578 (2015).</li><li>Richter, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structural+and+functional+analyses+of+TRP+ion+channels+in+the+photoreceptor+cells+of+Drosophila+melanogaster:+Dissertation+for+obtaining+the+academic+degree+Dr.+rer,+nat.">Structural and functional analyses of TRP ion channels in the photoreceptor cells of Drosophila melanogaster: Dissertation for obtaining the academic degree Dr. rer, nat.</a> University of Karlsruhe (TH). , (2007).</li><li>Gambis, A., Dourlen, P., Steller, H., Mollereau, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two-color+in+vivo+imaging+of+photoreceptor+apoptosis+and+development+in+Drosophila.">Two-color in vivo imaging of photoreceptor apoptosis and development in Drosophila.</a> Developmental Biology. 351 (1), 128-134 (2011).</li><li>Hardie, R. C., Liu, C. -. H., Randall, A. S., Sengupta, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+tracking+of+phosphoinositides+in+Drosophila+photoreceptors.">In vivo tracking of phosphoinositides in Drosophila photoreceptors.</a> Journal of Cell Science. 128 (23), 4328-4340 (2015).</li><li>Chakrabarti, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+dPIP5K+dependent+pool+of+phosphatidylinositol+4,5+bisphosphate+(PIP2)+is+required+for+G-protein+coupled+signal+transduction+in+Drosophila+photoreceptors.">A dPIP5K dependent pool of phosphatidylinositol 4,5 bisphosphate (PIP2) is required for G-protein coupled signal transduction in Drosophila photoreceptors.</a> PLoS Genetics. 11 (1), 1004948 (2015).</li></ol>

The authors have nothing to disclose.

The applicability of fluorescence proteins and simplicity of screening by DPP imaging and retinal water immersion microscopy has been proven to be successful by many groups12. Strategies similar to the ones presented here have been used in several genetic screens to detect defects in rhodopsin expression levels, homeostasis, retinal organization, or cellular integrity with the help of Rh1::eGFP17,18,19,20,21. As explained above, translocating fusion proteins like TRPL::eGFP may be used to detect whether protein trafficking processes from and to the rhabdomere are impaired, for example, in mutagenesis screens. On the other hand, permanently rhabdomeral fusion proteins like TRP::eGFP may be used to investigate rhabdomere morphology. DPP and water immersion microscopy are regarded as excellent methods for high-throughput analyses and screens within the model of the Drosophila eye. It is, however, recommended to ultimately perform (immuno)histochemical analyses of isolated ommatidia or cryo-sectioned retinal tissue to corroborate results obtained by DPP or water immersion microscopy. Paired with high-resolution fluorescence microscopy, these techniques allow interpretations toward precise subcellular localizations. In the following, a few guidelines are offered on how to troubleshoot the imaging processes of the DPP and retinal water immersion microscopy that is particularly important for correct quantitative analyses.
Concerning the DPP, if the superposition of fluorescence is blurry or indistinct, the settings may not be right. One possibility is that the eye is not facing the objective precisely radially or that the focus is set to the peripheral regions of the eye. Since the ommatidial layout of photoreceptor cells displays mirror symmetry at the dorsoventral midline, an unusual pattern in the DPP may be seen, if the signals are detected from both hemispheres of the eye. The quality of the DPP also heavily relies on a shallow depth of field to generate an optical section through the compound eye. If the surface of the flypad displays autofluorescence, the fly may be placed on a small piece of black non-fluorescent cardboard on top of the flypad. An inability to record a DPP may be connected to a variety of underlying causes, including suboptimal fly positioning, low expression levels of the fluorescent protein, a disorganized retinal architecture, or degenerative processes. When imaging the DPP, it should also be noted that cytoplasmic fluorescence in white-eyed flies may diffuse the borders of the signal from the rhabdomeres, and expression levels of protein may vary significantly between individuals and affect the quality of the resulting DPP. Thus, in the case of fluorescent DPP imaging, it is advantageous to use flies with pigmented eyes. Due to the absorption of fluorescence signals from the cell bodies, the resulting DPP may appear sharper than in white-eyed flies.
To avoid blurry images in the water immersion microscopy, image acquisition needs to be performed quickly after fly preparation to prevent reawakening and movements of the fly. For high-quality images, the brightest fluorescence signals from the eye have to almost reach saturation in all images if using white-eyed flies. This can be achieved by adjusting the exposure time during imaging. Since pigmentation hinders detection of cytosolic eGFP fluorescence, the described approach for white-eyed flies to adjust exposure time would always result in strong rhabdomeral fluorescence signals in pigmented eyes, distorting subsequent interpretation and quantification. Additionally, by using pigmented eyes, photobleaching due to strong exposure to blue light may lead to weaker signals and thus misinterpretation of data. Depending on the context (i.e., lethal&#160;or non-lethal), two alternative procedures are presented in the protocol for pigmented eyes. In general, the non-lethal variation is to be preferred with pigmented eyes, since the same fly can be measured repeatedly, and the assessment of a specific exposure time for each individual fly results in more precise quantifications and higher reproducibility than the determination of a mean exposure time generated from a sample. However, the non-lethal variation also demands more care in the handling of animals to ensure their survival during repeated measurements. This can be achieved by using only low pressured air to move flies into the pipette tip and carefully using tweezers to free the fly from the pipette tip after imaging. Since the flies are not submerged in ice-chilled water during the procedure, there is an increased risk of reawakening. As a result of the repeated measurements from individual specimens, the entire series of measurements is only valid if the fly stays unharmed from beginning to end. As a result, the non-lethal variation is very labor-intensive and time-consuming. The lethal variation, on the other hand, allows a higher throughput since multiple flies can be pinned in succession on the same insect pin and recorded sequentially in one image acquisition round. However, multiple flies on the same pin also require even quicker imaging to avoid reawakening and movements. A last disadvantage of the lethal variation is obviously the inability to recover the animals after imaging for subsequent applications (e.g., crosses, repeated measurements).
Regarding quantification of translocation, this Protocol presents formula (1), which considers fluorescence signals from the rhabdomere as well as the cell body and thus generates a very good estimate for the relative distribution of the translocating protein within the cell (Figure 12D). Due to the lack of fluorescence measurements from the cell body in the case of pigmented eyes, formula (2) is offered, which works well for this frequently encountered issue (Figure 13B). Aside from eye pigmentation, there are other circumstances that prevent the quantification with either of these formulae. One of these instances is presented in Figure 12A,B in the form of the lolattd12 mutant. In this mutant, specifically, the absence of photoreceptor cell R7 significantly affects the ommatidial morphology and thus the ability to acquire reasonable intensity measurements of background fluorescence from this region. If the same method of translocation quantification is being used despite these issues, the results display high amounts of variance, are difficult to compare with controls, and may potentially be misinterpreted. The example of the lolattd12 mutant thus illustrates the limits of this quantification method which inevitably relies on a &#34;wild type&#34;-like morphology for correct intensity measurements. In rare cases in which protein translocation defects are to be quantified in morphologically defective ommatidia, the Protocol presented here fails and has to be adapted accordingly. This includes appropriate changes to the formula in order to exclude measurements that cannot be obtained reasonably or to include additional measurements of other representative regions.
In the case of the quantification of degenerative processes, fluorescence intensity thresholds turned out to not be a good indicator, since there is too much variance within and between the rhabdomeres. This variance is averaged out over all 18 measured rhabdomeres in the method for quantifying translocation. However, due to the individual evaluation of each rhabdomere in the assessment of degeneration, the variance in fluorescence intensities cannot be averaged out. Furthermore, intensity is only one of the characteristics that need to be considered. Other important aspects are contrast and the sharpness of edges. Consequently, this protocol presents a quantification method based on categorizing the morphology of rhabdomeres according to their fluorescence by eye. This is necessarily subjective to some extent. Nevertheless, given enough experience, this protocol has proven a useful, fast, and reproducible method of quantification for degeneration (Figure 14) and has been published multiple times22,10. As with all subjective evaluations, it is always advisable to perform them in a blinded context. Protocols from other groups usually only use two categories to quantify retinal degeneration in Drosophila (presence or absence of rhabdomeres), which may be less subjective, but also less precise. Additionally, this way of quantification still has the issue of defining a cut-off point between the two categories, which may be equally hard to define objectively.
With regards to modifications and advanced applications of the methods presented here, it is also possible to assess the relative amount of two fluorescent fusion proteins simultaneously by double color imaging. For example, by using TRPL::HcRed&#160;and TRP::eGFP, TRPL::HcRed&#160;translocation has been assessed in consideration of rhabdomeral integrity in the same fly23. Double color imaging is also used in the Tomato/GFP-FLP/FRT method, e.g., to identify factors inducing apoptosis24. This variation makes it possible to visualize the degeneration of photoreceptor cells via ninaC::eGFP in fluorescently labeled cell clones marked by ninaC::tdTomato. Another application of water immersion microscopy is the monitoring of phosphoinositide turnover in photoreceptors. For this purpose, fluorescently tagged phosphoinositide-binding domains such as the pleckstrin homology (PH) domain from PLC&#948;1 are used. These fluorescence probes allow tracking of phosphoinositide degradation over time by detecting the decrease of rhabdomeral fluorescence caused by the diffusion of the probe from the rhabdomere to the cytosol25,26.

By delivering and removing proteins to and from the plasma membrane, membrane protein trafficking in neurons controls the plasma membrane equipment with receptors as well as ion channels and, as a result, regulates neuronal function. Misregulation or defects in protein trafficking typically have detrimental effects on cells and result in neuronal degeneration. In humans, this may cause neurodegenerative diseases such as Alzheimer&#39;s and Parkinson&#39;s disease or Retinitis pigmentosa1. Photoreceptors in the compound eye of Drosophila melanogaster have become an in vivo model system for studying membrane protein trafficking2.&#160;This is not only due to the genetic versatility of Drosophila that allows effective genetic screens, but also because all essential components of the light-absorbing photoreceptor membrane are characterized in great detail and efficient microscopic techniques are available that can be applied to the fly eye. These techniques are the focus of this article.
In Drosophila photoreceptor cells, the apical plasma membrane forms a densely packed stack of microvilli along one side of the cell, termed rhabdomere. The rhabdomeres of photoreceptor cells R1-6 are arranged in a characteristic trapezoidal pattern while photoreceptor cells R7 and R8 form a single rhabdomere in the center of this trapezoid3. Membrane protein trafficking is needed for a regulated turnover of rhabdomeral membrane proteins such as rhodopsin and the light-activated TRP (transient receptor potential) and TRPL (TRP-like) ion channels to assure the proper amount of these phototransduction proteins in the rhabdomere. Photoreceptor membrane proteins are synthesized in the endoplasmic reticulum and transported via the Golgi apparatus to the rhabdomere. Following activation of rhodopsin by light, a rhodopsin molecule can either become inactivated by absorption of a second photon or can be removed from the rhabdomere by clathrin-mediated endocytosis. Endocytosed rhodopsin either becomes degraded in the lysosome or is recycled back to the rhabdomere4,5. The ion channel TRPL is also internalized following activation of the phototransduction cascade and undergoes a light-dependent translocation between the rhabdomere (where it is located when flies are kept in the dark) and an ER-enriched storage compartment in the cell body (to which it is transported within several hours upon illumination)6,7,8,9,10. In contrast to endocytosed rhodopsin, only small amounts of TRPL are degraded via the endolysosomal pathway, and the majority is stored intracellularly instead and recycled back to the rhabdomere upon dark adaptation6. TRPL can thus be used for analyzing light-triggered trafficking of plasma membrane proteins. Drosophila photoreceptor cells are also employed for studying neuronal degeneration. Photoreceptor cell degeneration is frequently determined by assessing the structure of rhabdomeres, which disintegrate as a result of degenerative processes5.
In order to study the subcellular localization of TRPL and rhodopsin in photoreceptor cells or photoreceptor cell degeneration, two fluorescence microscopy methods that differ with respect to analysis speed and resolution have been applied here. A very fast, non-invasive method that can be used for genetic screens but with a limited spatial resolution is the detection of fluorescence in the deep pseudopupil (DPP). The DPP is an optical phenomenon of arthropod compound eyes whose geometric origin has been explained in detail by Franceschini and Kirschfeld in 197111. In short, on several optical planes below the retina overlay-images of rhabdomeres from adjacent ommatidia can be observed. On a focal plane through the center of the eye&#39;s curvature, these superimposed projections form an image that resembles the trapezoidal layout of rhabdomeres in a single ommatidium only orders of magnitude larger. This phenomenon can also be observed independently of exogenous expression of fluorescence proteins (e.g. TRPL::eGFP8), which nonetheless make the DPP easier to detect (Figure&#160;1A-A&#39;&#39;)12. A second non-invasive method is water immersion microscopy that relies on imaging fluorescently tagged proteins after optically neutralizing the eyes&#39; dioptric apparatus with water (Figure&#160;1B-C&#39;&#39;)12. Using the water immersion method, the relative amount of TRPL::eGFP in the rhabdomeres or cell body can be assessed quantitatively for individual photoreceptor cells. Furthermore, non-translocating fluorescence-tagged proteins can be utilized to evaluate rhabdomeral integrity and to determine the time course of a potential degeneration in a quantitative manner, as described here.
While recordings of the DPP are by far the easiest and fastest of these methods to perform, the spatial resolution of data they generate is limited. In addition, there are numerous reasons why a DPP may be absent, which are not necessarily discernible by DPP imaging itself. Since the DPP represents a summation of several ommatidia, information about individual cells is lost. Thus, low-resolution DPP imaging serves an important function in screening large numbers of flies but should generally be followed by higher resolution recordings by way of water immersion microscopy. Water immersion micrographs allow interpretations about individual cells, developmental defects, eye morphology, protein mislocalization or retinal degeneration as well as quantification of these effects. This Protocol describes these two techniques in detail.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63375/63375fig01.jpg" /> 
Figure 1: Overview of microscopy variations for the Drosophila eye presented in this Protocol. Schematic representations and exemplary micrographs of (A-A&#39;&#39;) fluorescent deep pseudopupil (DPP) imaging, (B-B&#39;&#39;) lethal water immersion microscopy of fluorescent rhabdomeres, and (C-C&#39;&#39;) non-lethal water drop microscopy of fluorescent rhabdomeres. Scale bar (A&#39;&#39;): 100 &#181;m. Scale bars (B&#39;&#39;-C&#39;&#39;): 10 &#181;m. The figure has been modified from reference13. <a href="https://www.jove.com/files/ftp_upload/63375/63375fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>15 mL centrifuge tube</td>
			<td>Greiner Bio-One</td>
			<td>188271</td>
			<td></td>
		</tr>
		<tr>
			<td>CO2 anaesthesia fly pad</td>
			<td>Flystuff</td>
			<td>59-172</td>
			<td></td>
		</tr>
		<tr>
			<td>Cold light lamp (KL 1500 LCD)</td>
			<td>Zeiss</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fiji/ImageJ</td>
			<td>NIH</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescence microscope with UV lamp, camera, filter set and software (AxioImager.Z1m, Axiocam 530 mono, 38 HE, ZEN2 blue edition)</td>
			<td>Zeiss</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescent tube (Lumilux T8, L 30W/840, 4000 K, G13) 
			[1750 Lux, Ee470nm = 298 &#181;W cm-2, Ee590nm = 215 &#181;W cm-2] 
			and [760 Lux, Ee470nm = 173 &#181;W cm-2, Ee590nm = 147 &#181;W cm-2]</td>
			<td>Osram</td>
			<td>4050300518039</td>
			<td></td>
		</tr>
		<tr>
			<td>Laboratory pipette (20-200 &#181;L)</td>
			<td>Eppendorf</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Object slide</td>
			<td>Roth</td>
			<td>0656.1</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dish (94 mm)</td>
			<td>Greiner Bio-One</td>
			<td>633102</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette tips (200 &#181;L)</td>
			<td>Labsolute</td>
			<td>7695844</td>
			<td></td>
		</tr>
		<tr>
			<td>Plasticine (Blu-Tack)</td>
			<td>Bostik</td>
			<td>30811745</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereo microscope (SMZ445)</td>
			<td>Nikon</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Stereo microscope with UV lamp, camera, filer set and software (MZ16F, MC170 HD, GFP3, LAS 4.12)</td>
			<td>Leica</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

studying membrane protein trafficking in drosophila photoreceptor cells using egfp-tagged proteins

Membrane protein trafficking regulates the incorporation and removal of receptors and ion channels into the plasma membrane. This process is fundamentally important for cell function and cell integrity of neurons. Drosophila photoreceptor cells have become a model for studying membrane protein trafficking. Besides rhodopsin, which upon illumination becomes internalized from the photoreceptor membrane and is degraded, the transient receptor potential-like (TRPL) ion channel in Drosophila exhibits a light-dependent translocation between the rhabdomeral photoreceptor membrane (where it is located in the dark) and the photoreceptor cell body (to which it is transported upon illumination). This intracellular transport of TRPL can be studied in a simple and non-invasive way by expressing eGFP-tagged TRPL in photoreceptor cells. The eGFP fluorescence can then be observed either in the deep pseudopupil or by water immersion microscopy. These methods allow detection of fluorescence in the intact eye and are therefore useful for high-throughput assays and genetic screens for Drosophila mutants defective in TRPL translocation. Here, the preparation of flies, the microscopic techniques, as well as quantification methods used to study this light-triggered translocation of TRPL are explained in detail. These methods can be applied also for trafficking studies on other Drosophila photoreceptor proteins, for example, rhodopsin. In addition, by using eGFP-tagged rhabdomeral proteins, these methods can be used to assess the degeneration of photoreceptor cells.

Transgenic Drosophila flies expressing a TRPL::eGFP fusion protein under the control of the rhodopsin 1 promoter have been generated. In these flies, TRPL::eGFP is expressed in photoreceptor cells R1-6 of the compound eye and displays an illumination-dependent localization. When flies are kept in the dark, TRPL::eGFP is incorporated into the outer rhabdomeres. After illumination for several hours, TRPL translocates into the cell body where it is stored in an ER-enriched compartment.8,10 When TRPL::eGFP is located in the rhabdomeres, its fluorescence can be observed in the DPP. However, rhabdomeral fluorescence disappears in the light as a result of translocation of TRPL::eGFP (Figure 11A,B). This has been used to perform a genetic screen for mutants defective in the internalization of TRPL::eGFP (Figure 11C,D)14,15. To allow isolation of potentially homozygous lethal mutations, this screen was carried out using the yeast Flp/FRT system for the generation of somatic cell clones in the developing eye tissue (i.e., mosaic eyes). Besides the simplicity of the DPP assay, other big advantages of this fluorescence detection method are its high throughput as well as its non-invasive character allowing identification of mutations in living flies and to use these flies for the generation of stable mutant fly stocks.
<img alt="Figure 11" class="xfigimg" src="/files/ftp_upload/63375/63375fig11v4.jpg" /> 
Figure 11: Representative results from a genetic screen on TRPL translocation defects. Male flies were mutagenized with ethyl methanesulfonate (EMS) and crossed to females expressing TRPL::eGFP in photoreceptor cells R1-6. The resulting F1-generation with homozygously mutant Flp/FRT-induced mosaic eyes was screened for defects in TRPL translocation by imaging fluorescent deep pseudopupils (DPP) after a period of dark adaptation (left column) as well as light adaptation (right column). (A,B) Non-mutated flies were carried along as control for regular light-induced TRPL translocation. The non-fluorescent DPP is indicated by an arrow. (C,D) An exemplary result of isolated mutants defective in light-triggered TRPL translocation. Scale bar: 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/63375/63375fig11v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
TRPL translocation defective (ttd) mutants that were isolated from this genetic screen have later been identified and their defects characterized in more detail with the help of water immersion imaging. Water immersion microscopy has been applied to assess the localization of proteins in the rhabdomeres and&#160;cell bodies&#160;or to track&#160;defects in the rhabdomeral structure due to photoreceptor degeneration quantitatively as outlined in the Protocol. Figure 12A,B illustrates quantification of the amount of TRPL::eGFP present in the rhabdomeres of flies with non-mutant and lolattd12 mutant mosaic eyes in the darkness, in light, and after a second dark-adaptation. In control (non-mutant) flies, TRPL::eGFP translocates out of the rhabdomeres into the cell body after 16 h of illumination resulting in a decrease of rhabdomeral TRPL::eGFP to about 45% as compared to the dark-adapted state. The second dark incubation raises the rhabdomeral fluorescence again to 95% of the initial value. The lolattd12 mutant was isolated by DPP screening due to its TRPL translocation defect. Accordingly, the TRPL::eGFP fluorescence pattern in water immersion images of rhabdomeres does not appear to change as drastically after 16 h of illumination and subsequent dark-adaption for 24 h. However, the images also reveal that lolattd12 mutants are developmentally defective, which is evidenced by the occasional absence of photoreceptor cells as described previously for other lola mutants16. These morphological defects result in an inability to measure a valid background signal, preventing a correct quantification using formula (1). In contrast, vps35MH20 is a mutant that displays a TRPL translocation defect without affecting the ommatidial morphology in this way10. In this mutant, the quantification method described here can detect a statistically highly significant recycling defect (Figure 12C,D).
<img alt="Figure 12" class="xfigimg" src="/files/ftp_upload/63375/63375fig12v2.jpg" /> 
Figure 12: Representative results of a TRPL translocating defect in lola and vps35 mutant flies and limits of the quantification method. (A) TRPL::eGFP-expressing non-mutant and lolattd12 mutant mosaic-eyed flies were incubated in the dark for 3 days, illuminated with orange light for 16 h, and dark-adapted for 24 h a second time. Images of fluorescent rhabdomeres were taken by way of water immersion microscopy to record subcellular TRPL::eGFP localization as described in step 3.2.6. (B) Quantification of rhabdomeral fluorescence of non-mutant (gray) and lolattd12 mutant (green) animals as described in step 3.3.9 using formula (1). Error bars represent the SEM. (C) TRPL::eGFP-expressing non-mutant and vps35MH20 mutant mosaic-eyed flies were incubated in the dark for 3 days, illuminated with orange light for 16 h, and dark-adapted for 24 h a second time. Images of fluorescent rhabdomeres were taken by way of water immersion microscopy to record subcellular TRPL::eGFP localization as described in step 3.2.6. (D) Quantification of rhabdomeral fluorescence of non-mutant (gray) and vps35MH20 mutant (red) animals as described in step 3.3.9 using formula (1). Error bars represent SEM. Statistical significance was calculated as multiple comparisons with Bonferroni correction after ANOVA (ns, not significant; *, p &#8804; 0.05; ***, p &#8804; 0.001). Scale bar: 10 &#181;m. Panels C and D have been modified from reference10. <a href="https://www.jove.com/files/ftp_upload/63375/63375fig12v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
As already mentioned in the Protocol, eye pigmentation significantly influences the resulting water immersion image. White-eyed flies allow the detection of fluorescence signals from both the rhabdomere and the cell body, thus enabling a fairly precise determination of the TRPL::eGFP distribution ratio. In contrast, in red-eyed flies TRPL::eGFP signal is only detectable in the rhabdomeres but not in the cell bodies&#160;(Figure 13A). This is due to pigment molecules absorbing the emitted light from TRPL::eGFP. Nevertheless, by using formula (2) for pigmented eyes, quantification nicely reveals TRPL::eGFP translocation behavior in white- as well as red-eyed flies. Accordingly, the corresponding values of rhabdomere fluorescence decrease from 100% in the dark-adapted state to 25% and 5% after illumination and increase again to 80% and 90% as a result of a second dark incubation (Figure 13B).
<img alt="Figure 13" class="xfigimg" src="/files/ftp_upload/63375/63375fig13v2.jpg" /> 
Figure 13: Representative results of TRPL::eGFP fluorescence in white- and red-eyed flies. (A) TRPL::eGFP-expressing white- and red-eyed flies were incubated in the dark for 1 day, illuminated with orange light for 16 h, and dark-adapted for 24 h a second time. Images of fluorescent rhabdomeres were taken by way of non-lethal water drop microscopy to record subcellular TRPL::eGFP localization as described in steps 3.2.6 (white-eyed flies) and 3.2.8 (red-eyed flies). (B) Quantification of rhabdomeral fluorescence in white-eyed (gray) and red-eyed (red) flies. White-eyed flies were quantified using formula (1) and red-eyed flies, using formula (2), as described in step 3.3.9. Error bars represent SEM. Scale bar: 10 &#181;m. <a href="https://www.jove.com/files/ftp_upload/63375/63375fig13v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
For assessing photoreceptor degeneration, a degeneration index based on the fluorescence of TRP::eGFP in the rhabdomeres can be determined (see Protocol). This quantification method is illustrated in Figure 10. In the representative results here, non-mutant and sdhAttd11 mutant mosaic-eyed flies were kept in a 12 h light / 12 h dark cycle for 2 weeks and the integrity of rhabdomeres was investigated every 2-4 days by observing TRP::eGFP using water immersion microscopy (Figure 14A). The resulting curve shows a decline of the degeneration index in the mutant but not in control flies (Figure 14B).
<img alt="Figure 14" class="xfigimg" src="/files/ftp_upload/63375/63375fig14v2.jpg" /> 
Figure 14: Representative results of light-induced retinal degeneration in sdhA mutant flies. (A) TRP::eGFP-expressing non-mutant and sdhAttd11 mutant mosaic-eyed flies were incubated for 14 days in a 12 h light / 12 h dark cycle at 25 &#176;C. Images of fluorescent rhabdomeres were taken by way of water immersion microscopy at regular time intervals to record retinal health. (B) Quantification of rhabdomeral fluorescence of wild type (gray) and sdhAttd11 mutant (orange) flies as described in step 3.4. Error bars represent SEM. Scale bar: 10 &#181;m. <a href="https://www.jove.com/files/ftp_upload/63375/63375fig14v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Studying Membrane Protein Trafficking in Drosophila Photoreceptor Cells Using eGFP-Tagged Proteins at JoVE.com

Studying Membrane Protein Trafficking in Drosophila Photoreceptor Cells Using eGFP-Tagged Proteins

Here, non-invasive methods are described for localization of photoreceptor membrane proteins and assessment of retinal degeneration in the Drosophila compound eye using eGFP fluorescence.

Studying Membrane Protein Trafficking in Drosophila Photoreceptor Cells Using eGFP-Tagged Proteins

Membrane protein trafficking regulates the incorporation and removal of receptors and ion channels into the plasma membrane. This process is fundamentally ...

studying-membrane-protein-trafficking-drosophila-photoreceptor-cells

Universidad San Sebastian

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JoVE Journal

Biology

2.7K Views.  University of Hohenheim. Here, non-invasive methods are described for localization of photoreceptor membrane proteins and assessment of retinal degeneration in the Drosophila compound eye using eGFP fluorescence.

Video: Studying Membrane Protein Trafficking in Drosophila Photoreceptor Cells Using eGFP-Tagged Proteins

Studying Aggression in Drosophila (fruit flies)

Aggression is an innate behavior that evolved in the framework of defending or obtaining resources. This complex social behavior is influenced by genetic, hormonal and environmental factors. In many organisms, aggression is critical to survival but controlling and suppressing aggression in distinct contexts also has become increasingly important. In recent years, invertebrates have become increasingly useful as model systems for investigating the genetic and systems biological basis of complex social behavior. This is in part due to the diverse repertoire of behaviors exhibited by these organisms. In the accompanying video, we outline a method for analyzing aggression in Drosophila whose design encompasses important eco-ethological constraints. Details include steps for: making a fighting chamber; isolating and painting flies; adding flies to the fight chamber; and video taping fights. This approach is currently being used to identify candidate genes important in aggression and in elaborating the neuronal circuitry that underlies the output of aggression and other social behaviors.

Studying Aggression in Drosophila fruit flies

Aggression is an innate behavior that evolved in the framework of defending or obtaining resources.  This complex social behavior is influenced by ...

Staining of Proteins in Gels with Coomassie G-250 without Organic Solvent and Acetic Acid

In classical protein staining protocols using Coomassie Brilliant Blue (CBB), solutions with high contents of toxic and flammable organic solvents (Methanol, Ethanol or 2-Propanol) and acetic acid are used for fixation, staining and destaining of proteins in a gel after SDS-PAGE. To speed up the procedure, heating the staining solution in the microwave oven for a short time is frequently used. This usually results in evaporation of toxic or hazardous Methanol, Ethanol or 2-Propanol and a strong smell of acetic acid in the lab which should be avoided due to safety considerations. In a protocol originally published in two patent applications by E.M. Wondrak (US2001046709 (A1), US6319720 (B1)), an alternative composition of the staining solution is described in which no organic solvent or acid is used. The CBB is dissolved in bidistilled water (60-80mg of CBB G-250 per liter) and 35 mM HCl is added as the only other compound in the staining solution. The CBB staning of the gel is done after SDS-PAGE and thorough washing of the gel in bidistilled water. By heating the gel during the washing and staining steps, the process can be finished faster and no toxic or hazardous compunds are evaporating. The staining of proteins occurs already within 1 minute after heating the gel in staining solution and is fully developed after 15-30 min with a slightly blue background that is destained completely by prolonged washing of the stained gel in bidistilled water, without affecting the stained protein bands.

A short protocol for protein staining with Coomassie Brilliant Blue (CBB) G-250 in polyacrylamide gels is described without using organic solvents or acetic acid as in the classical staining procedures with CBB.

In classical protein staining protocols using Coomassie Brilliant Blue (CBB), solutions with high contents of toxic and flammable organic solvents ...

Crystallizing Membrane Proteins for Structure Determination using Lipidic Mesophases

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the lipidic cubic phase or in meso method. The method has been shown to be quite versatile in that it has been used to solve X-ray crystallographic structures of prokaryotic and eukaryotic proteins, proteins that are monomeric, homo- and hetero-multimeric, chromophore-containing and chromophore-free, and alpha-helical and beta-barrel proteins. Recent successes using in meso crystallization are the human engineered beta2-adrenergic and adenosine A2a G protein-coupled receptors. Protocols are presented for reconstituting the membrane protein into the monoolein-based mesophase, and for setting up crystallizations in the manual mode. Additional steps in the overall process, such as crystal harvesting, are to be addressed in future video articles. The time required to prepare the protein-loaded mesophase and to set up a crystallization plate manually is about one hour.

Herein is described the procedure implemented in the Caffrey Membrane Structural and Functional Biology Group to set up manually crystallization trials of membrane proteins in lipidic mesophases.

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the ...

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle microtubules to prevent anaphase onset until the chromosomes are properly connected. Cells use this mechanism to prevent aneuploidy or genomic instability, and hence cancers and other human diseases like birth defects and Alzheimer's1. A number of the SAC components such as Mad1, Mad2, Bub1, BubR1, Bub3, Mps1, Zw10, Rod and Aurora B kinase have been identified and they are all kinetochore dynamic proteins2. Evidence suggests that the kinetochore is where the SAC signal is initiated. The SAC prime regulatory target is Cdc20. Cdc20 is one of the essential APC/C (Anaphase Promoting Complex or Cyclosome) activators3 and is also a kinetochore dynamic protein4-6. When activated, the SAC inhibits the activity of the APC/C to prevent the destruction of two key substrates, cyclin B and securin, thereby preventing the metaphase to anaphase transition7,8. Exactly how the SAC signal is initiated and assembled on the kinetochores and relayed onto the APC/C to inhibit its function still remains elusive.
Drosophila is an extremely tractable experimental system; a much simpler and better-understood organism compared to the human but one that shares fundamental processes in common. It is, perhaps, one of the best organisms to use for bio-imaging studies in living cells, especially for visualization of the mitotic events in space and time, as the early embryo goes through 13 rapid nuclear division cycles synchronously (8-10 minutes for each cycle at 25 &deg;C) and gradually organizes the nuclei in a single monolayer just underneath the cortex9.
Here, I present a bio-imaging method using transgenic Drosophila expressing GFP (Green Fluorescent Protein) or its variant-targeted proteins of interest and a Leica TCS SP2 confocal laser scanning microscope system to study the SAC function in flies, by showing images of GFP fusion proteins of some of the SAC components, Cdc20 and Mad2, as the example.

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The kinetochore is where the SAC initiates its signal monitoring the mitotic segregation of the sister chromatids. A method is described to visualize the recruitment and turnover of one of the kinetochore proteins and its coordination with the chromosome motion in Drosophila embryos using a Leica laser scanning confocal system.

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle ...

Use of a Robot for High-throughput Crystallization of Membrane Proteins in Lipidic Mesophases

Structure-function studies of membrane proteins greatly benefit from having available high-resolution 3-D structures of the type provided through macromolecular X-ray crystallography (MX). An essential ingredient of MX is a steady supply of ideally diffraction-quality crystals. The in meso or lipidic cubic phase (LCP) method for crystallizing membrane proteins is one of several methods available for crystallizing membrane proteins. It makes use of a bicontinuous mesophase in which to grow crystals. As a method, it has had some spectacular successes of late and has attracted much attention with many research groups now interested in using it. One of the challenges associated with the method is that the hosting mesophase is extremely viscous and sticky, reminiscent of a thick toothpaste. Thus, dispensing it manually in a reproducible manner in small volumes into crystallization wells requires skill, patience and a steady hand. A protocol for doing just that was developed in the Membrane Structural &amp; Functional Biology (MS&amp;FB) Group1-3. JoVE video articles describing the method are available1,4. 
The manual approach for setting up in meso trials has distinct advantages with specialty applications, such as crystal optimization and derivatization. It does however suffer from being a low throughput method. Here, we demonstrate a protocol for performing in meso crystallization trials robotically. A robot offers the advantages of speed, accuracy, precision, miniaturization and being able to work continuously for extended periods under what could be regarded as hostile conditions such as in the dark, in a reducing atmosphere or at low or high temperatures. An in meso robot, when used properly, can greatly improve the productivity of membrane protein structure and function research by facilitating crystallization which is one of the slow steps in the overall structure determination pipeline. 
In this video article, we demonstrate the use of three commercially available robots that can dispense the viscous and sticky mesophase integral to in meso crystallogenesis. The first robot was developed in the MS&amp;FB Group5,6. The other two have recently become available and are included here for completeness.	
An overview of the protocol covered in this article is presented in Figure 1. All manipulations were performed at room temperature (~20 &deg;C) under ambient conditions.

Herein is described a robotic approach to high-throughput crystallization of membrane proteins in lipidic mesophases for use in structure determination using macromolecular X-ray crystallography. Three robots capable of handling the viscous and sticky protein-laden mesophase integral to the method are introduced.

Structure-function studies of membrane proteins greatly benefit from having available high-resolution 3-D structures of the type provided through ...

Harvesting and Cryo-cooling Crystals of Membrane Proteins Grown in Lipidic Mesophases for Structure Determination by Macromolecular Crystallography

An important route to understanding how proteins function at a mechanistic level is to have the structure of the target protein available, ideally at atomic resolution. Presently, there is only one way to capture such information as applied to integral membrane proteins (Figure 1), and the complexes they form, and that method is macromolecular X-ray crystallography (MX). To do MX diffraction quality crystals are needed which, in the case of membrane proteins, do not form readily. A method for crystallizing membrane proteins that involves the use of lipidic mesophases, specifically the cubic and sponge phases1-5, has gained considerable attention of late due to the successes it has had in the G protein-coupled receptor field6-21 (<a href="http://www.mpdb.tcd.ie" target="_blank">www.mpdb.tcd.ie</a>). However, the method, henceforth referred to as the in meso or lipidic cubic phase method, comes with its own technical challenges. These arise, in part, due to the generally viscous and sticky nature of the lipidic mesophase in which the crystals, which are often micro-crystals, grow. Manipulating crystals becomes difficult as a result and particularly so during harvesting22,23. Problems arise too at the step that precedes harvesting which requires that the glass sandwich plates in which the crystals grow (Figure 2)24,25 are opened to expose the mesophase bolus, and the crystals therein, for harvesting, cryo-cooling and eventual X-ray diffraction data collection.
The cubic and sponge mesophase variants (Figure 3) from which crystals must be harvested have profoundly different rheologies4,26. The cubic phase is viscous and sticky akin to a thick toothpaste. By contrast, the sponge phase is more fluid with a distinct tendency to flow. Accordingly, different approaches for opening crystallization wells containing crystals growing in the cubic and the sponge phase are called for as indeed different methods are required for harvesting crystals from the two mesophase types. Protocols for doing just that have been refined and implemented in the Membrane Structural and Functional Biology (MS&amp;FB) Group, and are described in detail in this JoVE article (Figure 4). Examples are given of situations where crystals are successfully harvested and cryo-cooled. We also provide examples of cases where problems arise that lead to the irretrievable loss of crystals and describe how these problems can be avoided. In this article the Viewer is provided with step-by-step instructions for opening glass sandwich crystallization wells, for harvesting and for cryo-cooling crystals of membrane proteins growing in cubic and in sponge phases.

Herein is described procedures implemented in the Caffrey Membrane Structural and Functional Biology Group to harvest and cryo-cool membrane protein crystals grown in lipidic cubic and sponge phases for use in structure determination using macromolecular X-ray crystallography.

An important route to understanding how proteins function at a mechanistic level is to have the structure of the target protein available, ideally at ...

Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, we use the salivary glands of live mice since they provide several advantages. First, they can be easily exposed to enable access to the optics, and stabilized to facilitate the reduction of the motion artifacts due to heartbeat and respiration. This significantly facilitates imaging and tracking small subcellular structures. Second, most of the cell populations of the salivary glands are accessible from the surface of the organ. This permits the use of confocal microscopy that has a higher spatial resolution than other techniques that have been used for in vivo imaging, such as two-photon microscopy. Finally, salivary glands can be easily manipulated pharmacologically and genetically, thus providing a robust system to investigate biological processes at a molecular level.
In this study we focus on a protocol designed to follow the kinetics of the exocytosis of secretory granules in acinar cells and the dynamics of the apical plasma membrane where the secretory granules fuse upon stimulation of the beta-adrenergic receptors. Specifically, we used a transgenic mouse that co-expresses cytosolic GFP and a membrane-targeted peptide fused with the fluorescent protein tandem-Tomato. However, the procedures that we used to stabilize and image the salivary glands can be extended to other mouse models and coupled to other approaches to label in vivo cellular components, enabling the visualization of various subcellular structures, such as endosomes, lysosomes, mitochondria, and the actin cytoskeleton.

Intravital microscopy is a powerful tool that enables imaging various biological processes in live animals. In this article, we present a detailed method for imaging the dynamics of subcellular structures, such as the secretory granules, in the salivary glands of live mice.

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, ...

The Tomato/GFP-FLP/FRT Method for Live Imaging of Mosaic Adult Drosophila Photoreceptor Cells

The Drosophila eye is widely used as a model for studies of development and neuronal degeneration. With the powerful mitotic recombination technique, elegant genetic screens based on clonal analysis have led to the identification of signaling pathways involved in eye development and photoreceptor (PR) differentiation at larval stages. We describe here the Tomato/GFP-FLP/FRT method, which can be used for rapid clonal analysis in the eye of living adult Drosophila. Fluorescent photoreceptor cells are imaged with the cornea neutralization technique, on retinas with mosaic clones generated by flipase-mediated recombination. This method has several major advantages over classical histological sectioning of the retina: it can be used for high-throughput screening and has proved an effective method for identifying the factors regulating PR survival and function. It can be used for kinetic analyses of PR degeneration in the same living animal over several weeks, to demonstrate the requirement for specific genes for PR survival or function in the adult fly. This method is also useful for addressing cell autonomy issues in developmental mutants, such as those in which the establishment of planar cell polarity is affected.

The Tomato/GFP-FLP/FRT Method for Live Imaging of Mosaic Adult Drosophila Photoreceptor Cells

The Tomato/GFP-FLP/FRT method involves visualizing mosaic photoreceptor cells in living Drosophila. It can be used to follow individual photoreceptor cell fates in the retina for days or weeks. This method is ideal for studies of retinal degeneration and neurodegenerative diseases or photoreceptor cell development.

The Drosophila eye is widely used as a model  for studies of development and neuronal degeneration. With the powerful mitotic  recombination technique, ...

Easy Measurement of Diffusion Coefficients of EGFP-tagged Plasma Membrane Proteins Using k-Space Image Correlation Spectroscopy

Lateral diffusion and compartmentalization of plasma membrane proteins are tightly regulated in cells and thus, studying these processes will reveal new insights to plasma membrane protein function and regulation. Recently, k-Space Image Correlation Spectroscopy (kICS)1&#160;was developed to enable routine measurements of diffusion coefficients directly from images of fluorescently tagged plasma membrane proteins, that avoided systematic biases introduced by probe photophysics. Although the theoretical basis for the analysis is complex, the method can be implemented by nonexperts using a freely available code to measure diffusion coefficients of proteins. kICS calculates a time correlation function from a fluorescence microscopy image stack after Fourier transformation of each image to reciprocal (k-) space. Subsequently, circular averaging, natural logarithm transform and linear fits to the correlation function yields the diffusion coefficient. This paper provides a step-by-step guide to the image analysis and measurement of diffusion coefficients via kICS.
First, a high frame rate image sequence of a fluorescently labeled plasma membrane protein is acquired using a fluorescence microscope. Then, a region of interest (ROI) avoiding intracellular organelles, moving vesicles or protruding membrane regions is selected. The ROI stack is imported into a freely available code and several defined parameters (see Method section) are set for kICS analysis. The program then generates a &#34;slope of slopes&#34; plot from the k-space time correlation functions, and the diffusion coefficient is calculated from the slope of the plot. Below is a step-by-step kICS procedure to measure the diffusion coefficient of a membrane protein using the renal water channel aquaporin-3 tagged with EGFP as a canonical example.

This paper provides a step by step guide to the fluctuation analysis technique k-Space Image Correlation Spectroscopy (kICS) for measuring diffusion coefficients of fluorescently labeled plasma membrane proteins in live mammalian cells.

Lateral diffusion and compartmentalization of plasma membrane proteins are tightly regulated in cells and thus, studying these processes will reveal new ...

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and the inherent instability of many membrane proteins once solubilized in detergents. A protocol is described that combines ligation independent cloning of membrane proteins as GFP fusions with expression in Escherichia coli detected by GFP fluorescence. This enables the construction and expression screening of multiple membrane protein/variants to identify candidates suitable for further investment of time and effort. The GFP reporter is used in a primary screen of expression by visualizing GFP fluorescence following SDS polyacrylamide gel electrophoresis (SDS-PAGE). Membrane proteins that show both a high expression level with minimum degradation as indicated by the absence of free GFP, are selected for a secondary screen. These constructs are scaled and a total membrane fraction prepared and solubilized in four different detergents. Following ultracentrifugation to remove detergent-insoluble material, lysates are analyzed by fluorescence detection size exclusion chromatography (FSEC). Monitoring the size exclusion profile by GFP fluorescence provides information about the mono-dispersity and integrity of the membrane proteins in different detergents. Protein: detergent combinations that elute with a symmetrical peak with little or no free GFP and minimum aggregation are candidates for subsequent purification. Using the above methodology, the heterologous expression in E. coli of SED (shape, elongation, division, and sporulation) proteins from 47 different species of bacteria was analyzed. These proteins typically have ten transmembrane domains and are essential for cell division. The results show that the production of the SEDs orthologues in E. coli was highly variable with respect to the expression levels and integrity of the GFP fusion proteins. The experiment identified a subset for further investigation.

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

A streamlined approach to screening for the expression of recombinant membrane proteins in Escherichia coli based on fusion to green fluorescent protein is presented.

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and ...