Work in the Januschke laboratory is supported by Wellcome trust grants 100031/Z/12/A and 1000032/Z/12/A. The tissue imaging facility is supported by the grant WT101468 from Wellcome.

<ol><li>Restrepo, S., Zartman, J. J., Basler, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cultivation+and+live+imaging+of+Drosophila+imaginal+discs%5BTitle%5D)+AND+%22Methods+in+Molecular+Biology%22%5BJournal%5D)">Cultivation and live imaging of Drosophila imaginal discs.</a> Methods in Molecular Biology. 11478, 203-213 (2016).</li>
<li>Tsao, C. K., Ku, H. Y., Lee, Y. M., Huang, Y. F., Sun, Y. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Long+term+ex+vivo+culture+and+live+imaging+of+Drosophila+larval+imaginal+discs%5BTitle%5D)+AND+%22PloS+One%22%5BJournal%5D)">Long term ex vivo culture and live imaging of Drosophila larval imaginal discs.</a> PloS One. 11 (9), 0163744(2016).</li>
<li>Cabernard, C., Doe, C. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Apical%2Fbasal+spindle+orientation+is+required+for+neuroblast+homeostasis+and+neuronal+differentiation+in+Drosophila%5BTitle%5D)+AND+%22Developmental+Cell%22%5BJournal%5D)">Apical/basal spindle orientation is required for neuroblast homeostasis and neuronal differentiation in Drosophila.</a> Developmental Cell. 17 (1), 134-141 (2009).</li>
<li>Basto, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Flies+without+centrioles%5BTitle%5D)+AND+%22Cell%22%5BJournal%5D)">Flies without centrioles.</a> Cell. 125 (7), 1375-1386 (2006).</li>
<li>Januschke, J., Gonzalez, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+interphase+microtubule+aster+is+a+determinant+of+asymmetric+division+orientation+in+Drosophila+neuroblasts%5BTitle%5D)+AND+%22The+Journal+of+Cell+Biology%22%5BJournal%5D)">The interphase microtubule aster is a determinant of asymmetric division orientation in Drosophila neuroblasts.</a> The Journal of Cell Biology. 188 (5), 693-706 (2010).</li>
<li>Rusan, N. M., Peifer, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+role+for+a+novel+centrosome+cycle+in+asymmetric+cell+division%5BTitle%5D)+AND+%22Journal+of+Cell+Biology%22%5BJournal%5D)">A role for a novel centrosome cycle in asymmetric cell division.</a> Journal of Cell Biology. 177 (1), 13-20 (2007).</li>
<li>Dai, W., Montell, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Live+imaging+of+border+cell+migration+in+Drosophila%5BTitle%5D)+AND+%22Methods+in+Molecular+Biology%22%5BJournal%5D)">Live imaging of border cell migration in Drosophila.</a> Methods in Molecular Biology. 1407, 153-168 (2016).</li>
<li>Haigo, S. L., Bilder, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Global+tissue+revolutions+in+a+morphogenetic+movement+controlling+elongation%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">Global tissue revolutions in a morphogenetic movement controlling elongation.</a> Science. 331 (6020), 1071-1074 (2011).</li>
<li>Tissot, N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Distinct+molecular+cues+ensure+a+robust+microtubule-dependent+nuclear+positioning+in+the+Drosophila+oocyte%5BTitle%5D)+AND+%22Nature+Communications%22%5BJournal%5D)">Distinct molecular cues ensure a robust microtubule-dependent nuclear positioning in the Drosophila oocyte.</a> Nature Communications. 8, 15168(2017).</li>
<li>Martin, J. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Long-term+live+imaging+of+the+Drosophila+adult+midgut+reveals+real-time+dynamics+of+division%2C+differentiation+and+loss%5BTitle%5D)+AND+%22eLife%22%5BJournal%5D)">Long-term live imaging of the Drosophila adult midgut reveals real-time dynamics of division, differentiation and loss.</a> eLife. , (2018).</li>
<li>Cabernard, C., Doe, C. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Live+imaging+of+neuroblast+lineages+within+intact+larval+brains+in+Drosophila%5BTitle%5D)+AND+%22Cold+Spring+Harbor+Protocols%22%5BJournal%5D)">Live imaging of neuroblast lineages within intact larval brains in Drosophila.</a> Cold Spring Harbor Protocols. 2013 (10), 970-977 (2013).</li>
<li>Sabado, V., Nagoshi, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Single-cell+resolution+fluorescence+live+imaging+of+Drosophila+circadian+clocks+in+larval+brain+culture%5BTitle%5D)+AND+%22Journal+of+Visualized+Experiments%3A+JoVE%22%5BJournal%5D)">Single-cell resolution fluorescence live imaging of Drosophila circadian clocks in larval brain culture.</a> Journal of Visualized Experiments: JoVE. (131), (2018).</li>
<li>Lerit, D. A., Plevock, K. M., Rusan, N. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Live+imaging+of+Drosophila+larval+neuroblasts%5BTitle%5D)+AND+%22Journal+of+Visualized+Experiments%3A+JoVE%22%5BJournal%5D)">Live imaging of Drosophila larval neuroblasts.</a> Journal of Visualized Experiments: JoVE. (89), (2014).</li>
<li>Egger, B., van Giesen, L., Moraru, M., Sprecher, S. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((In+vitro+imaging+of+primary+neural+cell+culture+from+Drosophila%5BTitle%5D)+AND+%22Nature+Protocols%22%5BJournal%5D)">In vitro imaging of primary neural cell culture from Drosophila.</a> Nature Protocols. 8 (5), 958-965 (2013).</li>
<li>Homem, C. C., Reichardt, I., Berger, C., Lendl, T., Knoblich, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Long-term+live+cell+imaging+and+automated+4D+analysis+of+drosophila+neuroblast+lineages%5BTitle%5D)+AND+%22PloS+One%22%5BJournal%5D)">Long-term live cell imaging and automated 4D analysis of drosophila neuroblast lineages.</a> PloS One. 8 (11), 79588(2013).</li>
<li>Forer, A., Pickett-Heaps, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cytochalasin+D+and+latrunculin+affect+chromosome+behaviour+during+meiosis+in+crane-fly+spermatocytes%5BTitle%5D)+AND+%22Chromosome+Research%3A+An+iIternational+Journal+On+the+Molecular%2C+Supramolecular+and+Evolutionary+Aspects+of+Chromosome+Biology%22%5BJournal%5D)">Cytochalasin D and latrunculin affect chromosome behaviour during meiosis in crane-fly spermatocytes.</a> Chromosome Research: An iIternational Journal On the Molecular, Supramolecular and Evolutionary Aspects of Chromosome Biology. 6 (7), 533-549 (1998).</li>
<li>Pampalona, J., Januschke, J., Sampaio, P., Gonzalez, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Time-lapse+recording+of+centrosomes+and+other+organelles+in+Drosophila+neuroblasts%5BTitle%5D)+AND+%22Methods+in+Cell+Biology%22%5BJournal%5D)">Time-lapse recording of centrosomes and other organelles in Drosophila neuroblasts.</a> Methods in Cell Biology. 129, 301-315 (2015).</li>
<li>Ramat, A., Hannaford, M., Januschke, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Maintenance+of+miranda+localization+in+Drosophila+neuroblasts+involves+interaction+with+the+cognate+mRNA%5BTitle%5D)+AND+%22Current+Biology%3A+CB%22%5BJournal%5D)">Maintenance of miranda localization in Drosophila neuroblasts involves interaction with the cognate mRNA.</a> Current Biology: CB. 27 (14), 2101-2111 (2017).</li>
<li>Rebollo, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Functionally+unequal+centrosomes+drive+spindle+orientation+in+asymmetrically+dividing+drosophila+neural+stem+cells%5BTitle%5D)+AND+%22Developmental+Cell%22%5BJournal%5D)">Functionally unequal centrosomes drive spindle orientation in asymmetrically dividing drosophila neural stem cells.</a> Developmental Cell. 12 (3), 467-474 (2007).</li>
<li>Hannaford, M. R., Ramat, A., Loyer, N., Januschke, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((aPKC-mediated+displacement+and+actomyosin-mediated+retention+polarize+Miranda+in+Drosophila+neuroblasts%5BTitle%5D)+AND+%22eLife%22%5BJournal%5D)">aPKC-mediated displacement and actomyosin-mediated retention polarize Miranda in Drosophila neuroblasts.</a> eLife. 7, 166(2018).</li>
<li>Hannaford, M., Loyer, N., Tonelli, F., Zoltner, M., Januschke, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+chemical-genetics+approach+to+study+the+role+of+atypical+Protein+Kinase+C+in+Drosophila%5BTitle%5D)+AND+%22Development%22%5BJournal%5D)">A chemical-genetics approach to study the role of atypical Protein Kinase C in Drosophila.</a> Development. 146 (2), (2019).</li>
<li>Loyer, N., Januschke, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+last-born+daughter+cell+contributes+to+division+orientation+of+Drosophila+larval+neuroblasts%5BTitle%5D)+AND+%22Nature+Communications%22%5BJournal%5D)">The last-born daughter cell contributes to division orientation of Drosophila larval neuroblasts.</a> Nature Communications. 9 (1), 3745(2018).</li>
<li>Wilcockson, S. G., Ashe, H. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Drosophila+ovarian+germline+stem+cell+cytocensor+projections+dynamically+receive+and+attenuate+BMP+signaling%5BTitle%5D)+AND+%22Developmental+Cell%22%5BJournal%5D)">Drosophila ovarian germline stem cell cytocensor projections dynamically receive and attenuate BMP signaling.</a> Developmental Cell. 50 (3), 296-312 (2019).</li>
<li>Schneider, C. A., Rasband, W. S., Eliceiri, K. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((NIH+Image+to+ImageJ%3A+25+years+of+image+analysis%5BTitle%5D)+AND+%22Nature+methods%22%5BJournal%5D)">NIH Image to ImageJ: 25 years of image analysis.</a> Nature methods. 9 (7), 671-675 (2012).</li>
<li>Thevenaz, P., Ruttimann, U. E., Unser, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+pyramid+approach+to+subpixel+registration+based+on+intensity%5BTitle%5D)+AND+%22IEEE+Transactions+on+Image+Processing%22%5BJournal%5D)">A pyramid approach to subpixel registration based on intensity.</a> IEEE Transactions on Image Processing. 7 (1), 27-41 (1998).</li>
<li>Busse, B. MultiStackRef [software]. , Available from: 
 <a target="_blank" href="http://bradbusse.net/sciencedownloads.html">http://bradbusse.net/sciencedownloads.html</a> (2011).</li>
<li>Prasad, M., Montell, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cellular+and+molecular+mechanisms+of+border+cell+migration+analyzed+using+time-lapse+live-cell+imaging%5BTitle%5D)+AND+%22Developmental+Cell%22%5BJournal%5D)">Cellular and molecular mechanisms of border cell migration analyzed using time-lapse live-cell imaging.</a> Developmental Cell. 12 (6), 997-1005 (2007).</li>
<li>Chernysh, I. N., Nagaswami, C., Purohit, P. K., Weisel, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Fibrin+clots+are+equilibrium+polymers+that+can+be+remodeled+without+proteolytic+digestion%5BTitle%5D)+AND+%22Scientific+Reports%22%5BJournal%5D)">Fibrin clots are equilibrium polymers that can be remodeled without proteolytic digestion.</a> Scientific Reports. 2, 879(2012).</li>
<li>Liss, V., Barlag, B., Nietschke, M., Hensel, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Self-labelling+enzymes+as+universal+tags+for+fluorescence+microscopy%2C+super-resolution+microscopy+and+electron+microscopy%5BTitle%5D)+AND+%22Scientific+Reports%22%5BJournal%5D)">Self-labelling enzymes as universal tags for fluorescence microscopy, super-resolution microscopy and electron microscopy.</a> Scientific Reports. 5, 17740(2015).</li>
<li>Loyer, N. ImageJ macros [software]. , Available from: 
 <a target="_blank" href="https://github.com/nicolasloyer?tab=repositories">https://github.com/nicolasloyer?tab=repositories</a> (2020).</li>
</ol>

The authors have nothing to disclose.

Live imaging of whole mount D. melanogaster larval brains provides the opportunity to observe asymmetric neural stem cell divisions in conditions close to a physiological context. The first part of our protocol introduces our approach on the dissection of larval brains. As already stated13, a critical aspect of the preparation is to avoid damaging the brain. The most challenging aspect of this is to separate the brain from the neighboring imaginal disks without pulling excessively on the brain. Our approach to &#8220;cutting without pulling&#8221; is to perform either a grinding movement with the tips of one pair of forceps, or to slide a forceps tips along two other forceps tips holding the connection between tissues (Figure 1A) whereas Lerit et al. describe saw-like movements with a dissecting pin. We advise the experimenter to try all approaches and to adopt the best-suited one for themself. We also suggest using BSA- or FCS-coated pipette tips rather than dissection tools to transfer isolated brains. The coating prevents tissues from sticking to the plastic and allows the safe transfer of tissues while always keeping them completely immersed, without subjecting them to a possible transient deformation as they stick to the dissection tool during the transfer. Coated tips have other advantages to allow the transfer of several brains at once, which is particularly useful when it is critical to control the residency time of the samples within a particular medium; they are suitable for the transfer of other tissues, even fragile ones as, for example, fat bodies; they can accommodate a wide range of different sample sizes by using larger tips or cutting the extremity of the tip.
Next, this protocol describes the use of Fibrinogen clotting to immobilize larval brains on the coverslip of a culture dish. A brain is oriented within a drop of culture medium + Fibrinogen, after which clotting is induced by the addition of Thrombin. Fibrin formation is gradual, providing a time window during which the orientation of the sample can be fine-tuned, if necessary (Figure 2C). If a slight compression of the sample is required - which we advise against in the case of larval brains &#8211; it can also be finely adjusted by pressing the clot more or less close to the sample during steps 3.4.4 and 3.5.2. The main advantage of this Fibrinogen clotting over the protocol described in Lerit et al., in which brains are mounted between a gas permeable membrane and a coverslip, is the ability to replace the culture medium during live imaging. A possible advantage of using a gas permeable membrane over our protocol could be that it provides an optimal oxygenation of the samples, which could be more limited with our approach as brains are separated from the air by the clot and some medium. For this reason, although we did not assess the effect of the amount of culture medium within the culture dish, we suggest limiting the amount of culture medium on top of the clots while keeping the clots completely immersed.
As the manipulation of the clots can be experimentally difficult and time-consuming, we recommend that the experimenter first practices manipulating clots without samples. Modulating the volume of the drop of culture medium + Fibrinogen on the coverslip, the concentration of Fibrinogen or the volume and concentration of Thrombin could help making the preparation easier and could be important when adapting the protocol to other types of tissues. With enough experience manipulating the clots, several brains can be immobilized in one clot if necessary, although it complicates the fine-tuning of the orientation. Several clots can be formed on the coverslip, allowing, for example, to image samples of different genotypes. It is also possible to expose different clots on the same coverslip to different culture media, although care has to be taken never to mix the drops of culture medium covering the different clots. Once larval brains are immobilized and ready for live imaging, we advise to follow the recommendations of Lerit et al.13 to avoid excessive photodamage. We only add to these recommendations to not only maintain the brains at 25 &#176;C during live imaging with a stage incubator, but also to preheat this stage for at least 30 min before the start of imaging. In our hands, failing to do so systematically results in a strong focus drift.
It can be advantageous in some contexts to be able to perform correlated microscopy and fix and immunostain a sample after live imaging. However, a limitation of using Fibrin clots is that it is nearly impossible to mechanically isolate a brain from a clot without damaging it. A possible way to overcome this limitation could be to develop methods to degrade Fibrin clots with Plasmin or to induce clot disassembly by addition of a peptide mimicking the knobs allowing Fibrin polymerization28. We typically use Fibrin clots on samples ranging from single cells to 200 &#181;M long tissues, but we could also successfully immobilize in clots and image brains from adult bumblebees over 3 mm wide. The upper limit of the sample size that can be immobilized in clots remains to be determined. Similarly, although we usually image immobilized samples for 4 to 5 h, we successfully imaged neuroblasts in primary culture for up to 3 days, indicating that the clot at least does not interfere with longer term viability. On the contrary, clots can facilitate the regular replacement of the medium, a requirement for longer term imaging, without the need for devices such as peristaltic pumps for this purpose. While we have not measured the permeability parameters of fibrin clots, the fibrous gel-like nature of the clots appears to be penetrable by cell permeable molecules. We found that Latrunculin A, Colcemid or 1-NAPP1, HALO-tag ligands and other standard dyes used in cell biology reach the cells in the fibrin clot without any problems and have so far not encountered molecules that would be retained by the clot, which, however, is a possibility that needs to be empirically tested.
In conclusion, using Fibrin clots provide a reliable and relatively easy to implement a way of immobilizing living tissues for live imaging. Beyond its usefulness in limiting lateral drifting, the ability to change the culture medium during live imaging has proven invaluable for our chemical genetics approach to studying the asymmetric cell division of D. melanogaster neuroblasts21. We anticipate that this technique will be beneficial to studies in a wide range of different tissues, particularly if they involve chemical genetics or, for example, protein self-labeling29.
Our MultiHyperStackReg macro relies on the TurboReg ImageJ plugin, which aligns successive frames or slices of a stack, and the MultiStackReg ImageJ plugin, which allows to apply the transformations to other stacks. MultiHyperStackReg simply allows to apply these transformations to hyperstacks (stacks with at least four dimensions). As the use of MultiHyperStackReg, as we describe it, requires a lot of actions and can get quite time consuming, we also wrote the AutoHyperStackReg macro, which automatically performs all the steps described in steps 6.2 and 6.3. However, contrary to MultiHyperStackReg, AutoHyperStackReg currently lacks the possibility to calculate the alignment of a stack based on a subregion of this stack, an option that we found is sometimes crucial for a satisfying alignment. Another limitation of AutoHyperStackReg is that its use is restricted to translations and not the other transformations proposed by TurboReg and MultiStackReg, whereas MultiHyperStackReg can apply to a hyperstack any transformation type should the user need it. Future releases will implement these functions and will be available on GitHub30. Finally, our rotating linescan macro provides an easy way to quickly measure cortical signals in time lapses, even if the cortex changes its position or orientation over time. Whether its tracking mode or its non-tracking mode is best suited for analysis depends on the quality and consistency of the cortical signal. The tracking mode is easier to use as the user does not need to consider where the cortex will be for every timepoint and can accommodate large changes of position and orientation over time. However, it is only suitable if the cortical signal remains strong enough for detection during the entire movie: a failure to detect anything else than the cortex (e.g., a bright cytoplasmic compartment close to the cortex) can compromise the detection for every following timepoint (e.g., the cytoplasmic compartment moves away from the cortex and the cortical linescan follows it for the rest of the time lapse). The non-tracking mode does not have this pitfall as the detection is limited to the line originally drawn by the user (e.g., a bright cytoplasmic compartment may cause the detection to fail for a few timepoints, but as the cytoplasmic compartment moves away from the detection zone, proper detection of the cortex resumes), but does not behave well if the cortex position or orientation changes a lot over time. The next feature (which will be available on GitHub30) to be developed for the tracking mode will be the option to only analyze specified timepoints and to define a reference timepoint (rather than the first timepoint) before which and after which the detection will be performed, which will allow the user to at least avoid problematic timepoints.

Drosophila continues to be an important model system to study a vast range of biological questions and has excelled in advancing knowledge in many disciplines for decades. One feature that makes it particularly stand out is that it is especially well-suited for live imaging. The ability to monitor subcellular processes or the behavior of cells and tissues in real time continues to contribute to generating key concepts relevant to cell and developmental biology. Many successful protocols have been developed by different groups to maintain such Drosophila samples alive and healthy to study the behavior of the specimen and fluorescent molecules live. Organs and tissues from the fly such as imaginal discs1,2, the larval brain3,4,5,6, or egg chambers of adult females7,8,9, or the adult intestine10 for instance can be extracted and kept in culture for imaging with time-lapse microscopy. A key challenge for live imaging is to ensure that the specimen is kept close to the imaging surface for optimal resolution and immobile without compromising sample integrity that can be sensitive to mechanical impacts. To study neural stem cells, called neuroblasts or neurons in the Drosophila larval brain, for instance, keeping samples close to the imaging surface can be achieved by covering them with culture media in sealed imaging chambers11,12,13. However, this approach does not easily allow media exchange thus limiting experimental room for maneuver. Alternatively, samples can be prepared and placed on coverslips treated to increase adhesion of cells and allow multi-point visiting microscopy and extended live-cell imaging in open culture dishes, that potentially allow media exchange, but do not provide easy means to prevent sample drift or loss during media exchange14,15.
The Fibrin clot method originally developed to study meiosis in living crane-fly spermatocytes16 is specifically suited to overcome these problems. Fibrinogen is dissolved in relevant culture medium, the samples placed and oriented in a drop of this solution on an adequate glass-surface imaging chamber before Thrombin is added, resulting in the rapid formation of an insoluble Fibrin clot, a sticky fibrous mesh that adheres to the glass surface and the samples while ensuring access of the sample to the culture medium. The medium can then be exchanged without perturbing the clot or sample position. Culture medium exchange during imaging is, for instance, desirable when inhibitors are to be added as in the original method or for wash out or pulse chase experiments or when fluorescent dyes are to be added during live cell imaging while keeping cells and tissues in place. Fibrin clots are suitable for imaging of Drosophila tissues as well as individual cells13,17. Fibrin clots can further be used to help orient samples, that due to their shape or other properties would normally not be oriented in the desired way by modulating sample position within the Fibrin clot and the shape of the clot itself.
Here we provide an update on our current Fibrin clot-based imaging methods and provide tools for segmentation-based image analysis and focus on the use of this method to study neuroblast divisions in the developing fly brain. We routinely use the Fibrin clot method to follow multiple samples in different clots in the same culture dish. This allows i) imaging of subcellular events such as centrosome behavior or mRNA localization live18,19, ii) monitoring the behavior of samples upon pharmacological treatment of different genotypes under the same imaging settings using multi-point visiting microscopy20, iii) studying the effects of acute inhibition of enzymes21 and iv) minimizing drifting to study changes in the orientation of cell division of cells within their physiological environment upon targeted laser-ablation22. While unwanted effects of Thrombin and Fibrinogen and the Fibrin clot need to be empirically tested for each sample, this method is in principle suitable to immobilize any type of sample that is to be analyzed by live cell imaging and in Drosophila has been successfully used to study multiple aspects of neuroblasts biology5,19,20, but also the dynamics of cytosensor projections of female germ line stem cells23 and changes in polarity upon acute aPKC inhibition of follicle cells of Drosophila female egg chambers21.
Time-lapse fluorescent imaging results in the generation of complex time-resolved 3D data sets that require methods to extract this quantitative information. Here we describe our further development of ImageJ-based24 macros that can be used to correct for drifting in hyperstacks and custom-made ImageJ macros that allow semi-automated quantification of multi-channel fluorescence at the cell cortex developed to quantify cortical proteins in neuroblasts of Drosophila larval brains or of individual neuroblasts in primary cell culture.

<table><tbody><tr><td>Albumin bovine serum, BSA</td><td>Merck</td><td>5470</td><td></td></tr><tr><td>D-(+)-Glucose</td><td>Sigma</td><td>G7021</td><td></td></tr><tr><td>Dimethyl sulfoxide</td><td>Sigma</td><td>D2650</td><td></td></tr><tr><td>Dumont #55 Forceps</td><td>Fine Science Tools</td><td>11255-20</td><td></td></tr><tr><td>Fetal Bovie Serum, suitable for cell culture</td><td>Sigma</td><td>F7524</td><td></td></tr><tr><td>Fibrinogen from human plasma</td><td>Sigma</td><td>F3879</td><td></td></tr><tr><td>FluoroDish Cell Culture Dish - 35mm, 23 mm well</td><td>World Precision Instruments</td><td>FD35-100</td><td></td></tr><tr><td>PP1 Analog (1NA-PP1)</td><td>Merck Millipore</td><td>529579</td><td></td></tr><tr><td>Pyrex spot plate with nine depressions</td><td>Sigma</td><td>CLS722085-18EA</td><td></td></tr><tr><td>Schneider&#039;s Insect Medium</td><td>Sigma</td><td>S0146</td><td></td></tr><tr><td>Thrombin from bovine plasma</td><td>Merck</td><td>T4648</td><td></td></tr></tbody></table>

applications of immobilization of drosophila tissues with fibrin clots for live imaging

Drosophila is an important model system to study a vast range of biological questions. Various organs and tissues from different developmental stages of the fly such as imaginal discs, the larval brain or egg chambers of adult females or the adult intestine can be extracted and kept in culture for imaging with time-lapse microscopy, providing valuable insights into cell and developmental biology. Here, we describe in detail our current protocol for the dissection of Drosophila larval brains, and then present our current approach for immobilizing and orienting larval brains and other tissues on a glass coverslip using Fibrin clots. This immobilization method only requires the addition of Fibrinogen and Thrombin to the culture medium. It is suitable for high-resolution time lapse imaging on inverted microscopes of multiple samples in the same culture dish, minimizes the lateral drifting frequently caused by movements of the microscope stage in multi-point visiting microscopy and allows for the addition and removal of reagents during the course of imaging. We also present custom-made macros that we routinely use to correct for drifting and to extract and process specific quantitative information from time-lapse analysis.

Immobilization of Drosophila tissues with Fibrin clots and culture medium exchange during live imaging. 
After being dissected following the procedure presented in step 2 (Figure 1) and immobilized in Fibrin clots on the same coverslip following the procedure presented in step 3 (Figure 2), larval brains expressing GFP-tagged Bazooka (Baz::GFP), the fly ortholog of the polarity protein Par-3, were imaged for 30 min in time-lapse multi-position confocal imaging. Baz::GFP is combined with apkcas4, an allele that allows acute inhibition of atypical protein kinase C (aPKC) through the addition of the small ATP analog 1-NAPP120. 1-NAPP1 was therefore added to the culture medium following the procedure presented in step 4 and live imaging was resumed for 2.5 h. Control brains carrying a wild-type version of the kinase aPKC did not react to the ATP analog, whereas brains carrying additionally an analog-sensitive mutation of aPKC (apkcas4) displayed a contraction of the neuroepithelium and bright clusters of Baz in neuroblasts and their progeny (Figure 6, Video 1). Neuroblasts keep on dividing throughout the time-lapse, indicating that the tissue remains healthy, and brains show little drift despite the culture medium change, indicating that they are well immobilized. Similarly, after being dissected following the procedure presented in27, ovarioles expressing Baz::GFP were immobilized in Fibrin clots on the same coverslip and imaged for 8 min, after which application of 1-NAPP1 induced the contraction of apkcas4 mutant follicular cells but not of controls (Figure 7, Video 2).
Correction of lateral and focus drift using MultiHyperstackReg. 
A hyperstack displaying both lateral and focus drift (Video 3, left panel) was first corrected for lateral drift (step 6.2, Figure 3, Video 3, middle panel), then focus drift (step 6.3, Figure 4, Video 3, right panel) using the MultiStackReg plugin and the MultiHyperstackReg macro, resulting in a substantial reduction of the movements observed in the original hyperstack.
Cortical signal measurement using rotating linescans 
A neuroblast expressing Baz::GFP (green) and the transmembrane protein CD4 tagged with an infrared fluorescent protein (CD4::mIFP, red) was imaged during one mitosis. The CD4::mIFP signal was used as a reference to detect the cortex at various parts of the neuroblast with linescans (step 7, Figure 5, Figure 8A&#8211;C) and the Baz::GFP signal was measured at the detected positions Figure 8D). During their division, neuroblasts transiently polarized by establishing distinct and opposite cortical domains: the apical pole and the basal pole. Baz defines the apical pole and, consistently, the Baz::GFP signal increased at the apical pole of the neuroblast and decreased at the basal pole during the division (Figure 8C,D). The position of the cortex was satisfyingly tracked throughout the time-lapse (Figure 8C, Video 4). The Drosophila lines and genotypes are referenced in Supplemental Table 1&#8211;2.
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/61954/61954fig06.jpg" /> 
Figure 6: Application of an ATP analog on immobilized larval brains during live imaging.&#160;(A) Live imaging of two larval brains expressing Baz::GFP immobilized on the same coverslip. The culture medium is changed to a concentration of 10 &#181;M of the ATP analog 1-NAPP1 at 0 min. Control brains do not visibly react to the ATP analog whereas brains carrying an analog sensitive mutation of aPKC (aPKCAS4) display a contraction of the neuroepithelium (arrowheads) and bright clusters of Baz in neuroblasts and their progeny. Maximal intensity projection of 33 slices for a depth of 23 &#181;M. Scale bar: 20 &#181;M. (B) Magnification of the neuroepithelium. Scale bar: 10 &#181;M. <a href="https://www.jove.com/files/ftp_upload/61954/61954fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/61954/61954fig07.jpg" /> 
Figure 7: Application of an ATP analog on immobilized egg chambers during live imaging.&#160;Live imaging of two ovarioles expressing Baz::GFP immobilized on the same coverslip. The culture medium is changed to a concentration of 10 &#181;M of the ATP analog 1-NAPP1 at 0 min. Control egg chambers do not visibly react to the ATP analog whereas egg chambers carrying an analog sensitive mutation of aPKC (aPKCAS4) display a contraction of the epithelium (arrowheads) eventually leading to the apparent breakdown of adherens junctions. Maximal intensity projection of 33 slices for a depth of 27 &#181;M. Scale bar: 20 &#181;M. <a href="https://www.jove.com/files/ftp_upload/61954/61954fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/61954/61954fig08.jpg" /> 
Figure 8: Measurement of the cortical signal using rotating linescans.&#160;(A) Live D. melanogaster neuroblast expressing Baz::GFP (green) and CD4::mIFP (red). Cyan lines: initial scanning lines traced perpendicularly to various cortical zones to measure. Scale bar: 5 &#181;M. (B) Identification of the cortex (dark blue line) using cortical linescans and CD4::mIFP (red) as the reference channel. Cyan line: area defined by the &#8220;Measure around&#8221; parameter (here, 2 pixels), where the signal is measured after cortex detection. Scale bar: 2 &#181;m. (C) Cyan: cortical areas identified during a neuroblast division by the rotating linescans method from the initial scanning lines displayed in (A), using CD4::mIFP (red) as the reference channel. Scale bar: 5 &#181;M. Arrow: apical pole. Arrowhead: basal pole. (D) Measurement of the Baz::GFP signal intensity over time in the two corresponding zones identified by rotating linescans displayed in (C). During mitosis, Baz::GFP gets transiently enriched at the apical pole of the neuroblast and is depleted from the opposite basal pole. <a href="https://www.jove.com/files/ftp_upload/61954/61954fig08large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Video 1: Application of media exchange on immobilized larval brains during live imaging to add an inhibitor, presented in Figure 6, scale bar: 20 &#181;m. <a href="https://cloudflare.jove.com/files/ftp_upload/61954/MOV_1_Loyer_Januschke.mp4" target="_blank">Please click here to download this video.</a>
Video 2: Application of media exchange on immobilized egg chambers during live imaging to add an inhibitor, presented in Figure 7, scale bar: 20 &#181;m. <a href="https://cloudflare.jove.com/files/ftp_upload/61954/MOV_2_Loyer_Januschke.mp4" target="_blank">Please click here to download this video.</a>
Video 3: Correction of lateral and focus drift using MultiHyperStackReg. Live imaging of a neuroblast expressing the membrane probe PH::GFP. Xy: single focal plane. Xz: orthogonal reslice. Left: before correction of the drift. Middle: after correction of the lateral drift. Right: after correction the lateral and focus drift, scale bar: 10 &#181;m. <a href="https://cloudflare.jove.com/files/ftp_upload/61954/MOV_3_Loyer_Januschke.mp4" target="_blank">Please click here to download this video.</a>
Video 4: Detection of the cortex using rotating linescans, presented in Figure 8, scale bar: 5 &#181;m. <a href="https://cloudflare.jove.com/files/ftp_upload/61954/MOV_4_Loyer_Januschke.mp4" target="_blank">Please click here to download this video.</a>
Supplemental Table 1: Genotypes of imaged Drosophila tissues. <a href="https://cloudflare.jove.com/files/ftp_upload/61954/Supplemental_Table_1_genotypes.xlsx" target="_blank">Please click here to download this table.</a>
Supplemental Table 2: Origin of transgenes used. <a href="https://cloudflare.jove.com/files/ftp_upload/61954/Supplemental_Table_2_origine_of_transgenes.xlsx" target="_blank">Please click here to download this table.</a>
Supplemental Coding File 1:&#160;Origin of transgenes used.&#160;<a href="https://cloudflare.jove.com/files/ftp_upload/61954/Code1_MultiHyperstackReg.ijm" target="_blank">Please click here to download this file.</a>
Supplemental Coding File 2:&#160;Origin of transgenes used.&#160;<a href="https://cloudflare.jove.com/files/ftp_upload/61954/Code2_AutoHyperstackReg.ijm" target="_blank">Please click here to download this file.</a>
Supplemental Coding File 3:&#160;Origin of transgenes used.&#160;<a href="https://cloudflare.jove.com/files/ftp_upload/61954/Code3_RotatingLineScan.ijm" target="_blank">Please click here to download this file.</a>

Watch this Scientific Journal Video about Applications of Immobilization of Drosophila Tissues with Fibrin Clots for Live Imaging at JoVE.com

Applications of Immobilization of Drosophila Tissues with Fibrin Clots for Live Imaging

This protocol describes applications of sample immobilization using Fibrin clots, limiting drifting, and allowing the addition and washout of reagents during live imaging. Samples are transferred to a drop of Fibrinogen containing culture medium on the surface of a coverslip, after which polymerization is induced by adding thrombin.

Applications of Immobilization of Drosophila Tissues with Fibrin Clots for Live Imaging

Drosophila is an important model system to study a vast range of biological questions. Various organs and tissues from different developmental stages of ...

applications-immobilization-drosophila-tissues-with-fibrin-clots-for

Research

JoVE Journal

Developmental Biology

3.1K Views.  University of Dundee. Dit protocol beschrijft toepassingen van monsterimmobilisatie met behulp van Fibrine-stolsels, waardoor driften worden beperkt en de toevoeging en washout van reagentia tijdens live beeldvorming mogelijk is. Monsters worden overgebracht naar een druppel Fibrinogeen met kweekmedium op het oppervlak van een afdeklip, waarna polymerisatie wordt geïnduceerd door toevoeging van trombine.

Video: Applications of Immobilization of Drosophila Tissues with Fibrin Clots for Live Imaging

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of the Drosophila larva makes it an attractive model organism for live imaging studies. However, an important challenge for live imaging techniques is to noninvasively immobilize and position an animal on the microscope. This protocol presents a simple and easy to use method for immobilizing and imaging Drosophila larvae on a polydimethylsiloxane (PDMS) microfluidic device, which we call the 'larva chip'. The larva chip is comprised of a snug-fitting PDMS microchamber that is attached to a thin glass coverslip, which, upon application of a vacuum via a syringe, immobilizes the animal and brings ventral structures such as the nerve cord, segmental nerves, and body wall muscles, within close proximity to the coverslip. This allows for high-resolution imaging, and importantly, avoids the use of anesthetics and chemicals, which facilitates the study of a broad range of physiological processes. Since larvae recover easily from the immobilization, they can be readily subjected to multiple imaging sessions. This allows for longitudinal studies over time courses ranging from hours to days. This protocol describes step-by-step how to prepare the chip and how to utilize the chip for live imaging of neuronal events in 3rd instar larvae. These events include the rapid transport of organelles in axons, calcium responses to injury, and time-lapse studies of the trafficking of photo-convertible proteins over long distances and time scales. Another application of the chip is to study regenerative and degenerative responses to axonal injury, so the second part of this protocol describes a new and simple procedure for injuring axons within peripheral nerves by a segmental nerve crush.

Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

Drosophila larvae are an attractive model system for live imaging due to their translucent cuticle and powerful genetics. This protocol describes how to utilize a single-layer PDMS device, called the &#39;larva chip&#39; for live imaging of cellular processes within neurons of 3rd instar Drosophila larvae.

Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of ...

Bioengineering

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle synapses, we developed a larval tissue preparation that had features of live intact larvae, but also had good optical properties.  This new preparation also allowed for access of perfusates to the synapse.  We used fly larvae, immersed them in artificial hemolymph, and relaxed their normal rhythmic body contractions by chilling them. Next we dissected off the posterior segments of each animal and with a blunt insect pin pushed the mouth parts backward through the body cavity.  This everted the larval body wall, like turning a sock inside-out.  We completed the dissection with ultra-fine dissection scissors and thus exposed the visceral side of the body wall muscles. The glial structures at the NMJ expressed membrane targeted GFP under the control of glial specific promoters. The post-synaptic membrane, the SSR (Subsynaptic Reticula) in muscle expressed synaptically targeted dsRed. We needed to acutely label the motor neuron terminals, the third part of the synapse. To do this we applied primary antibodies to HRP, conjugated to a far-red emitting flurophore.  To test for dye diffusion properties into the perisynaptic space between the motor neuron terminals and the SSR, we applied a solution of large Dextran molecules conjugated to far-red emitting flurophore and collected images.

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

We described structural features of the Glia-neuromuscular synapses in a novel Inside-out tissue preparation of live fly larvae using fluorescent dyes with confocal microscopy. We labeled live neuron terminals with fluorescent primary antibodies to HRP, and also visualized the perisynaptic space with fluorescent Dextrans.

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle ...

Biology

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as tissue morphogenesis, cell-cell adhesion, or cell death. Drosophila embryos expressing GFP are readily imaged using either stereoscopic or confocal microscopy. A goal of any live-imaging protocol is to minimize detrimental effects such as dehydration and hypoxia. Previous protocols for preparing Drosophila embryos for live-imaging analysis have involved placing dechorionated embryos in halocarbon oil and sandwiching them between a halocarbon gas-permeable membrane and a coverslip1-3. The introduction of compression through mounting embryos in this manner represents an undesirable complication for any biomechanical-based analysis of morphogenesis. Our method, which we call the hanging drop protocol, results in excellent viability of embryos during live imaging and does not require that embryos be compressed. Briefly, the hanging drop protocol involves the placement of embryos in a drop of halocarbon oil that is suspended from a coverslip, which is, in turn, fixed in position over a humid chamber. In addition to providing gas exchange and preventing dehydration, this arrangement takes advantage of the buoyancy of embryos in halocarbon oil to prevent them from drifting out of position during timelapse acquisition. This video describes in detail how to collect and prepare Drosophila embryos for live imaging using the hanging drop protocol. This protocol is suitable for imaging dechorionated embryos using stereomicroscopy or any upright compound fluorescence microscope.

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A simple, inexpensive, and effective method of preparing Drosophila embryos for live-imaging analysis is presented. Our protocol provides humidity and gas exchange and does not compress the Drosophila embryo. This method is suitable for GFP-based live imaging of Drosophila embryos using a stereomicroscope or upright compound microscope.

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as ...

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we present a protocol for the mounting of Drosophila melanogaster embryos and subsequent live imaging of fluorescently labeled hemocytes, the embryonic macrophages of this organism. Using the Gal4-uas system1 we drive the expression of a variety of genetically encoded, fluorescently tagged markers in hemocytes to follow their developmental dispersal throughout the embryo. Following collection of embryos at the desired stage of development, the outer chorion is removed and the embryos are then mounted in halocarbon oil between a hydrophobic, gas-permeable membrane and a glass coverslip for live imaging. In addition to gross migratory parameters such as speed and directionality, higher resolution imaging coupled with the use of fluorescent reporters of F-actin and microtubules can provide more detailed information concerning the dynamics of these cytoskeletal components.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Drosophila hemocytes disperse over the entirety of the developing embryo. This protocol demonstrates how to mount and image these migrations using embryos with fluorescently labelled hemocytes.

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we ...

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three preparations of the brain and visual system that cover the range from the developing eye disc-brain complex in the developing pupae to individual eye and brain dissection from adult flies. All protocols are optimized for the live culture of the preparations. However, we also present the conditions for fixed tissue immunohistochemistry where applicable. Finally, we show live imaging conditions for these preparations using conventional and resonant 4D confocal live imaging in a perfusion chamber. Together, these protocols provide a basis for live imaging on different time scales ranging from functional intracellular assays on the scale of minutes to developmental or degenerative processes on the scale of many hours.

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

This protocol describes three Drosophila preparations: 1) adult brain dissection, 2) adult retina dissection and 3) developing eye disc- brain complexes dissection. Emphasis is laid on special preparation techniques and conditions for live imaging, although all preparations can be used for fixed tissue immunohistochemistry.

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three ...

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal lines have enabled biological processes to be studied in the context of a living, and in some instances even behaving, organism. In this protocol we will describe how to anesthetize intact Drosophila larvae, using the volatile anesthetic desflurane, to follow the development and plasticity of synaptic populations at sub-cellular resolution1-3. While other useful methods to anesthetize Drosophila melanogaster larvae have been previously described4,5,6,7,8, the protocol presented herein demonstrates significant improvements due to the following combined key features: (1) A very high degree of anesthetization; even the heart beat is arrested allowing for lateral resolution of up to 150 nm1, (2) a high survival rate of &gt; 90% per anesthetization cycle, permitting the recording of more than five time-points over a period of hours to days2 and (3) a high sensitivity enabling us in 2 instances to study the dynamics of proteins expressed at physiological levels. In detail, we were able to visualize the postsynaptic glutamate receptor subunit GluR-IIA expressed via the endogenous promoter1 in stable transgenic lines and the exon trap line FasII-GFP1. (4) In contrast to other methods4,7 the larvae can be imaged not only alive, but also intact (i.e. non-dissected) allowing observation to occur over a number of days1. The accompanying video details the function of individual parts of the in vivo imaging chamber2,3, the correct mounting of the larvae, the anesthetization procedure, how to re-identify specific positions within a larva and the safe removal of the larvae from the imaging chamber.

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

This protocol describes a reliable method for anesthetization and imaging of intact Drosophila melanogaster larvae. We have utilized the volatile anesthetic desflurane to allow for repetitive imaging at sub-cellular resolution and re-identification of structures for up to a few days1.

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. Single-molecule imaging is gaining momentum as it provides exceptional access to the visualization of individual molecules in living cells. Here, we describe a technique that we developed to perform single-particle tracking photo-activated localization microscopy (sptPALM) in Drosophila larvae. Synaptic communication relies on key presynaptic proteins that act by docking, priming, and promoting the fusion of neurotransmitter-containing vesicles with the plasma membrane. A range of protein-protein and protein-lipid interactions tightly regulates these processes and the presynaptic proteins therefore exhibit changes in mobility associated with each of these key events. Investigating how mobility of these proteins correlates with their physiological function in an intact live animal is essential to understanding their precise mechanism of action. Extracting protein mobility with high resolution in vivo requires overcoming limitations such as optical transparency, accessibility, and penetration depth. We describe how photoconvertible fluorescent proteins tagged to the presynaptic protein Syntaxin-1A can be visualized via slight oblique illumination and tracked at the motor nerve terminal or along the motor neuron axon of the third instar Drosophila larva.

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Here we illustrate how single molecule photo-activated localization microscopy can be carried out on the motor nerve terminal of a live Drosophila melanogaster third instar larva.

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. ...

Neuroscience

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

The circadian pacemaker circuit orchestrates rhythmic behavioral and physiological outputs coordinated with environmental cues, such as day/night cycles. The molecular clock within each pacemaker neuron generates circadian rhythms in gene expression, which underlie the rhythmic neuronal functions essential to the operation of the circuit. Investigation of the properties of the individual molecular oscillators in different subclasses of pacemaker neurons and their interaction with neuronal signaling yields a better understanding of the circadian pacemaker circuit. Here, we present a time-lapse fluorescent microscopy approach developed to monitor the molecular clockwork in clock neurons of cultured Drosophila larval brain. This method allows the multi-day recording of the rhythms of genetically encoded fluorescent circadian reporters at single-cell resolution. This setup can be combined with pharmacological manipulations to closely analyze real-time response of the molecular clock to various compounds. Beyond circadian rhythms, this multipurpose method in combination with powerful Drosophila genetic techniques offers the possibility to study diverse neuronal or molecular processes in live brain tissue.

Single-cell Resolution Fluorescence Live Imaging of Drosophila Circadian Clocks in Larval Brain Culture

The goal of this protocol is to establish ex vivo Drosophila larval brain culture optimized to monitor circadian molecular rhythms with long-term fluorescence time-lapse imaging. The application of this method to pharmacological assays is also discussed.

The circadian pacemaker circuit orchestrates rhythmic behavioral and physiological outputs coordinated with environmental cues, such as day/night cycles. ...

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

The Drosophila oocyte has been established as a versatile system for investigating fundamental questions such as cytoskeletal function, cell organization, and organelle structure and function. The availability of various GFP-tagged proteins means that many cellular processes can be monitored in living cells over the course of minutes or hours, and using this technique, processes such as RNP transport, epithelial morphogenesis, and tissue remodeling have been described in great detail in Drosophila oocytes1,2.
The ability to perform video imaging combined with a rich repertoire of mutants allows an enormous variety of genes and processes to be examined in incredible detail. One such example is the process of ooplasmic streaming, which initiates at mid-oogenesis3,4. This vigorous movement of cytoplasmic vesicles is microtubule and kinesin-dependent5 and provides a useful system for investigating cytoskeleton function at these stages.
Here I present a protocol for time lapse imaging of living oocytes using virtually any confocal microscopy setup.

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

A protocol for live imaging of GFP-tagged proteins or autofluorescent structures in individual Drosophila oocytes is described.

The  Drosophila oocyte has been established as a versatile system for investigating  fundamental questions such as cytoskeletal function, cell ...

LarvaSPA, A Method for Mounting Drosophila Larva for Long-Term Time-Lapse Imaging

Live imaging is a valuable approach for investigating cell biology questions. The Drosophila larva is particularly suited for in vivo live imaging because the larval body wall and most internal organs are transparent. However, continuous live imaging of intact Drosophila larvae for longer than 30 min has been challenging because it is difficult to noninvasively immobilizeimmobilizing larvae for a long time. Here we present a larval mounting method called LarvaSPA that allows for continuous imaging of live Drosophila larvae with high temporal and spatial resolution for longer than 10 hours. This method involves partially attaching larvae to the coverslip using a UV-reactive glue and additionally restraining larval movement using a polydimethylsiloxane (PDMS) block. This method is compatible with larvae at developmental stages from second instar to wandering third instar. We demonstrate applications of this method in studying dynamic processes of Drosophila somatosensory neurons, including dendrite growth and injury-induced dendrite degeneration. This method can also be applied to study many other cellular processes that happen near the larval body wall.

LarvaSPA, A Method for Mounting Drosophila Larva for Long-Term Time-Lapse Imaging

This protocol describes a method for mounting Drosophila larvae to achieve longer than 10 h of uninterrupted time-lapse imaging in intact live animals. This method can be used to image many biological processes close to the larval body wall.

Live imaging is a valuable approach for investigating cell biology questions. The Drosophila larva is particularly suited for in vivo live imaging because ...

Applications of Immobilization of Drosophila Tissues with Fibrin Clots for Live Imaging

Samenvatting

Meer video's verkennen