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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cultivation+and+live+imaging+of+Drosophila+imaginal+discs.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long+term+ex+vivo+culture+and+live+imaging+of+Drosophila+larval+imaginal+discs.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Apical/basal+spindle+orientation+is+required+for+neuroblast+homeostasis+and+neuronal+differentiation+in+Drosophila.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Flies+without+centrioles.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+interphase+microtubule+aster+is+a+determinant+of+asymmetric+division+orientation+in+Drosophila+neuroblasts.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+role+for+a+novel+centrosome+cycle+in+asymmetric+cell+division.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+imaging+of+border+cell+migration+in+Drosophila.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Global+tissue+revolutions+in+a+morphogenetic+movement+controlling+elongation.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinct+molecular+cues+ensure+a+robust+microtubule-dependent+nuclear+positioning+in+the+Drosophila+oocyte.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long-term+live+imaging+of+the+Drosophila+adult+midgut+reveals+real-time+dynamics+of+division,+differentiation+and+loss.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+imaging+of+neuroblast+lineages+within+intact+larval+brains+in+Drosophila.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-cell+resolution+fluorescence+live+imaging+of+Drosophila+circadian+clocks+in+larval+brain+culture.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+imaging+of+Drosophila+larval+neuroblasts.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+imaging+of+primary+neural+cell+culture+from+Drosophila.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Long-term+live+cell+imaging+and+automated+4D+analysis+of+drosophila+neuroblast+lineages.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cytochalasin+D+and+latrunculin+affect+chromosome+behaviour+during+meiosis+in+crane-fly+spermatocytes.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Time-lapse+recording+of+centrosomes+and+other+organelles+in+Drosophila+neuroblasts.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Maintenance+of+miranda+localization+in+Drosophila+neuroblasts+involves+interaction+with+the+cognate+mRNA.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functionally+unequal+centrosomes+drive+spindle+orientation+in+asymmetrically+dividing+drosophila+neural+stem+cells.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=aPKC-mediated+displacement+and+actomyosin-mediated+retention+polarize+Miranda+in+Drosophila+neuroblasts.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+chemical-genetics+approach+to+study+the+role+of+atypical+Protein+Kinase+C+in+Drosophila.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+last-born+daughter+cell+contributes+to+division+orientation+of+Drosophila+larval+neuroblasts.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+ovarian+germline+stem+cell+cytocensor+projections+dynamically+receive+and+attenuate+BMP+signaling.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NIH+Image+to+ImageJ:+25+years+of+image+analysis.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+pyramid+approach+to+subpixel+registration+based+on+intensity.">A pyramid approach to subpixel registration based on intensity.</a> IEEE Transactions on Image Processing. 7 (1), 27-41 (1998).</li><li>Prasad, M., Montell, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cellular+and+molecular+mechanisms+of+border+cell+migration+analyzed+using+time-lapse+live-cell+imaging.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fibrin+clots+are+equilibrium+polymers+that+can+be+remodeled+without+proteolytic+digestion.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Self-labelling+enzymes+as+universal+tags+for+fluorescence+microscopy,+super-resolution+microscopy+and+electron+microscopy.">Self-labelling enzymes as universal tags for fluorescence microscopy, super-resolution microscopy and electron microscopy.</a> Scientific Reports. 5, 17740 (2015).</li><li>. 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&#39;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>

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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 ...

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JoVE Journal

Developmental Biology

3.0K Views.  University of Dundee. 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.

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

Live-imaging of the Drosophila Pupal Eye

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track during live cell imaging. These processes include: retinal rotation, cell growth and organismal movement. Additionally, irregularities in the topology of the epithelium, including subtle bumps and folds often introduced as the pupa is prepared for imaging, make it challenging to acquire in-focus images of more than a few ommatidia in a single focal plane. The workflow outlined here remedies these issues, allowing easy analysis of cellular processes during Drosophila pupal eye development. Appropriately-staged pupae are arranged in an imaging rig that can be easily assembled in most laboratories. Ubiquitin-DE-Cadherin:GFP and GMR-GAL4&#8211;driven UAS-&#945;-catenin:GFP are used to visualize cell boundaries in the eye epithelium 1-3. After deconvolution is applied to fluorescent images captured at multiple focal planes, maximum projection images are generated for each time point and enhanced using image editing software. Alignment algorithms are used to quickly stabilize superfluous motion, making individual cell behavior easier to track.

Live-imaging of the Drosophila Pupal Eye

This protocol presents an efficient method for imaging the live Drosophila pupal eye neuroepithelium. This method compensates for tissue movement and uneven topology, enhances visualization of cell boundaries through the use of multiple GFP-tagged junction proteins, and uses an easily-assembled imaging rig.

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track ...

In Vitro Differentiation of Mature Myofibers for Live Imaging

Skeletal muscles are composed of myofibers, the biggest cells in the mammalian body and one of the few syncytia. How the complex and evolutionarily conserved structures that compose it are assembled remains under investigation. Their size and physiological features often constrain manipulation and imaging applications. The culture of immortalized cell lines is widely used, but it can only replicate the early steps of differentiation.
Here, we describe a protocol that enables easy genetic manipulation of myofibers originating from primary mouse myoblasts. After one week of differentiation, the myofibers display contractility, aligned sarcomeres and triads, as well as peripheral nuclei. The entire differentiation process can be followed by live imaging or immunofluorescence. This system combines the advantages of the existing ex vivo and in vitro protocols. The possibility of easy and efficient transfection as well as the ease of access to all differentiation stages broadens the potential applications. Myofibers can subsequently be used not only to address relevant developmental and cell biology questions, but also to reproduce muscle disease phenotypes for clinical applications.

In Vitro Differentiation of Mature Myofibers for Live Imaging

Muscle cells are among the most complex eukaryotic cells. We present a protocol for the in vitro differentiation of highly mature myofibers that allows for genetic manipulation and clear imaging during all developmental stages.

Skeletal muscles are composed of myofibers, the biggest cells in the mammalian body and one of the few syncytia. How the complex and evolutionarily ...

Live Confocal Imaging of Developing Arabidopsis Flowers

The study of plant growth and development has long relied on experimental techniques using dead, fixed tissues and lacking proper cellular resolution. Recent advances in confocal microscopy, combined with the development of numerous fluorophores, have overcome these issues and opened the possibility to study the expression of several genes simultaneously, with a good cellular resolution, in live samples. Live confocal imaging provides plant biologists with a powerful tool to study development, and has been extensively used to study root growth and the formation of lateral organs on the flanks of the shoot apical meristem. However, it has not been widely applied to the study of flower development, in part due to challenges that are specific to imaging flowers, such as the sepals that grow over the flower meristem, and filter out the fluorescence from underlying tissues. Here, we present a detailed protocol to perform live confocal imaging on live, developing Arabidopsis flower buds, using either an upright or an inverted microscope.

Live confocal imaging provides biologists with a powerful tool to study development. Here, we present a detailed protocol for the live confocal imaging of developing Arabidopsis flowers.

The study of plant growth and development has long relied on experimental techniques using dead, fixed tissues and lacking proper cellular resolution. ...

Long-term Live Imaging of Drosophila Eye Disc

Live imaging provides the ability to continuously track dynamic cellular and developmental processes in real time. Drosophila larval imaginal discs have been used to study many biological processes, such as cell proliferation, differentiation, growth, migration, apoptosis, competition, cell-cell signaling, and compartmental boundary formation. However, methods for the long-term ex vivo culture and live imaging of the imaginal discs have not been satisfactory, despite many efforts. Recently, we developed a method for the long-term ex vivo culture and live imaging of imaginal discs for up to 18 h. In addition to using a high insulin concentration in the culture medium, a low-melting agarose was also used to embed the disc to prevent it from drifting during the imaging period. This report uses the eye-antennal discs as an example. Photoreceptor R3/4-specific m&#948;0.5-Ga4 expression was followed to demonstrate that photoreceptor differentiation and ommatidial rotation can be observed during a 10 h live imaging period. This is a detailed protocol describing this simple method.

Long-term Live Imaging of Drosophila Eye Disc

This protocol describes a detailed method for the long-term ex vivo culture and live imaging of a Drosophila imaginal disc. It demonstrates photoreceptor differentiation and ommatidial rotation within the 10 h period of live imaging of the eye disc. The protocol is simple and does not require expensive setup.

Live imaging provides the ability to continuously track dynamic cellular and developmental processes in real time. Drosophila larval imaginal discs have ...

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; for example, how the zygote polarizes to establish the body axis and how the embryo specifies various cell fates during organ formation. This manuscript describes an in vitro ovule culture method to perform live-cell imaging of developing zygotes and embryos of Arabidopsis thaliana. The optimized cultivation medium allows zygotes or early embryos to grow into fertile plants. By combining it with a poly(dimethylsiloxane) (PDMS) micropillar array device, the ovule is held in the liquid medium in the same position. This fixation is crucial to observe the same ovule under a microscope for several days from the zygotic division to the late embryo stage. The resulting live-cell imaging can be used to monitor the real-time dynamics of zygote polarization, such as nuclear migration and cytoskeleton rearrangement, and also the cell division timing and cell fate specification during embryo patterning. Furthermore, this ovule cultivation system can be combined with inhibitor treatments to analyze the effects of various factors on embryo development, and with optical manipulations such as laser disruption to examine the role of cell-cell communication.

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

This manuscript describes an in vitro ovule cultivation method that enables live-cell imaging of Arabidopsis zygotes and embryos. This method is utilized to visualize the intracellular dynamics during zygote polarization and the cell fate specification in developing embryos.

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; ...

Live Imaging of Mouse Secondary Palate Fusion

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to cleft secondary palate, a common congenital anomaly in humans. Secondary palate fusion has been extensively studied leading to several proposed cellular mechanisms that may mediate this process. However, these studies have been mostly performed on fixed embryonic tissues at progressive timepoints during development or in fixed explant cultures analyzed at static timepoints. Static analysis is limited for the analysis of dynamic morphogenetic processes such a palate fusion and what types of dynamic cellular behaviors mediate palatal fusion is incompletely understood. Here we describe a protocol for live imaging of ex vivo secondary palate fusion in mouse embryos. To examine cellular behaviors of palate fusion, epithelial-specific Keratin14-cre was used to label palate epithelial cells in ROSA26-mTmGflox reporter embryos. To visualize filamentous actin, Lifeact-mRFPruby reporter mice were used. Live imaging of secondary palate fusion was performed by dissecting recently-adhered secondary palatal shelves of embryonic day (E) 14.5 stage embryos and culturing in agarose-containing media on a glass bottom dish to enable imaging with an inverted confocal microscope. Using this method, we have detected a variety of novel cellular behaviors during secondary palate fusion. An appreciation of how distinct cell behaviors are coordinated in space and time greatly contributes to our understanding of this dynamic morphogenetic process. This protocol can be applied to mutant mouse lines, or cultures treated with pharmacological inhibitors to further advance understanding of how secondary palate fusion is controlled.

Here, we present a protocol for live imaging of mouse secondary palate fusion using confocal microscopy. This protocol can be used in combination with a variety of fluorescent reporter mouse lines, and with pathway inhibitors for mechanistic insight. This protocol can be adapted for live imaging in other developmental systems.

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to ...

Methods for Imaging Intracellular pH of the Follicle Stem Cell Lineage in Live Drosophila Ovarian Tissue

Changes in intracellular pH (pHi) play important roles in the regulation of many cellular functions, including metabolism, proliferation, and differentiation. Typically, pHi dynamics are determined in cultured cells, which are amenable to measuring and experimentally manipulating pHi. However, the recent development of new tools and methodologies has made it possible to study pHi dynamics within intact, live tissue. For Drosophila research, one important development was the generation of a transgenic line carrying a pHi biosensor, mCherry::pHluorin. Here, we describe a protocol that we routinely use for imaging live Drosophila ovarioles to measure pHi in the epithelial follicle stem cell (FSC) lineage in mCherry::pHluorin transgenic wild type lines; however, the methods described here can be easily adapted for other tissues, including the wing discs and eye epithelium. We describe techniques for expressing mCherry::pHluorin in the FSC lineage, maintaining ovarian tissue during live imaging, and acquiring and analyzing images to obtain pHi values.

Methods for Imaging Intracellular pH of the Follicle Stem Cell Lineage in Live Drosophila Ovarian Tissue

We provide a protocol for imaging intracellular pH of an epithelial stem cell lineage in live Drosophila ovarian tissue. We describe methods to generate transgenic flies expressing a pH biosensor, mCherry::pHluorin, image the biosensor using quantitative fluorescence imaging, generate standard curves, and convert fluorescence intensity values to pH values.

Changes in intracellular pH (pHi) play important roles in the regulation of many cellular functions, including metabolism, proliferation, and ...

Live Cell Imaging of Chromosome Segregation During Mitosis

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled by multiple mechanisms that include the Spindle Assembly Checkpoint (SAC). The SAC is part of a complex feedback system that is responsible for prevention of a cell progress through mitosis unless all chromosomal kinetochores have attached to spindle microtubules. Chromosomal lagging and abnormal chromosome segregation is an indicator of dysfunctional cell cycle control checkpoints and can be used to measure the genomic stability of dividing cells. Deregulation of the SAC can result in the transformation of a normal cell into a malignant cell through the accumulation of errors during chromosomal segregation. Implementation of the SAC and the formation of the kinetochore complex are tightly regulated by interactions between kinases and phosphatase such as Protein Phosphatase 2A (PP2A). This protocol describes live cell imaging of lagging chromosomes in mouse embryonic fibroblasts isolated from mice that had a knockout of the PP2A-B56&#947; regulatory subunit. This method overcomes the shortcomings of other cell cycle control imaging techniques such as flow cytometry or immunocytochemistry that only provide a snapshot of a cell cytokinesis status, instead of a dynamic spatiotemporal visualization of chromosomes during mitosis.

This protocol describes an easy and convenient method to label and visualize live chromosomes in mitotic cells using Histone2B-GFP BacMam 2.0 label and a spinning disc confocal microscopy system.

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled ...

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Coordinated muscle contractions are a form of rhythmic behavior seen early during development in Drosophila embryos. Neuronal sensory feedback circuits are required to control this behavior. Failure to produce the rhythmic pattern of contractions can be indicative of neurological abnormalities. We previously found that defects in protein O-mannosylation, a posttranslational protein modification, affect the axon morphology of sensory neurons and result in abnormal coordinated muscle contractions in embryos. Here, we present a relatively simple method for recording and analyzing the pattern of peristaltic muscle contractions by live imaging of late stage embryos up to the point of hatching, which we used to characterize the muscle contraction phenotype of protein O-mannosyltransferase mutants. Data obtained from these recordings can be used to analyze muscle contraction waves, including frequency, direction of propagation and relative amplitude of muscle contractions at different body segments. We have also examined body posture and taken advantage of a fluorescent marker expressed specifically in muscles to accurately determine the position of the embryo midline. A similar approach can also be utilized to study various other behaviors during development, such as embryo rolling and hatching.

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Here, we present a method to record embryonic muscle contractions in Drosophila embryos in a non-invasive and detail-oriented manner.

Coordinated muscle contractions are a form of rhythmic behavior seen early during development in Drosophila embryos. Neuronal sensory feedback circuits ...

Imaging Intranuclear Actin Rods in Live Heat Stressed Drosophila Embryos

The purpose of this protocol is to visualize intranuclear actin rods that assemble in live Drosophila melanogaster embryos following heat stress. Actin rods are a hallmark of a conserved, inducible Actin Stress Response (ASR) that accompanies human pathologies, including neurodegenerative disease. Previously, we showed that the ASR contributes to morphogenesis failures and reduced viability of developing embryos. This protocol allows the continued study of mechanisms underlying actin rod assembly and the ASR in a model system that is highly amenable to imaging, genetics and biochemistry. Embryos are collected and mounted on a coverslip to prepare them for injection. Rhodamine-conjugated globular actin (G-actinRed) is diluted and loaded into a microneedle. A single injection is made into the center of each embryo. After injection, embryos are incubated at elevated temperature and intranuclear actin rods are then visualized by confocal microscopy. Fluorescence recovery after photobleaching (FRAP) experiments may be performed on the actin rods; and other actin-rich structures in the cytoplasm can also be imaged. We find that G-actinRed polymerizes like endogenous G-actin and does not, on its own, interfere with normal embryo development. One limitation of this protocol is that care must be taken during injection to avoid serious injury to the embryo. However, with practice, injecting G-actinRed into Drosophila embryos is a fast and reliable way to visualize actin rods and can easily be used with flies of any genotype or with the introduction of other cellular stresses, including hypoxia and oxidative stress.

Imaging Intranuclear Actin Rods in Live Heat Stressed Drosophila Embryos

The goal of this protocol is to inject Rhodamine-conjugated globular actin into Drosophila embryos and image intranuclear actin rod assembly following heat stress.

The purpose of this protocol is to visualize intranuclear actin rods that assemble in live Drosophila melanogaster embryos following heat stress. Actin ...

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

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Live-imaging of the Drosophila Pupal Eye

In Vitro Differentiation of Mature Myofibers for Live Imaging

Live Confocal Imaging of Developing Arabidopsis Flowers

Long-term Live Imaging of Drosophila Eye Disc

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

Live Imaging of Mouse Secondary Palate Fusion

Methods for Imaging Intracellular pH of the Follicle Stem Cell Lineage in Live Drosophila Ovarian Tissue

Live Cell Imaging of Chromosome Segregation During Mitosis

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Imaging Intranuclear Actin Rods in Live Heat Stressed Drosophila Embryos