Biz Andreas Schönle, Biyofiziksel Kimya, Almanya ve David J. Sandström, Ulusal Ruh Sağlığı Enstitüsü, Sağlık, Bethesda, MD, ABD Ulusal Sağlık Enstitüleri teknik danışmanlık için Max-Plank Enstitüsü teşekkür ederim. Biz Kötting Frank, Avrupa Nörobilim Enstitüsü, Göttingen görüntüleme odasının inşa için ve anesthetization cihazı teşekkür ederiz. Bu çalışma, Alzheimer Forschung TMRYZ Girişimi hibeleri tarafından desteklenen, Çin Burs Kurulu, SBH Hücresel ve Moleküler Nörobilim, Tübingen Üniversitesi Graduate School of bir burs ile bir dostluk tarafından desteklenen oldu.

A) Meclis görüntüleme odası <ol><li> Larva seçilmiş evre (yaklaşık 24 saatlik bir gözlem aralığı bırakarak 25 ° C örneğin sahne dolaşıp kadar erken 3. instar larva) seçin. </li><li> Larva, kültür ortamı ve dab kuru temiz su ile durulayın. </li><li> Halokarbon yağı ince bir film tabakası ile odasının alt elemanı, larva yüz bölgenin merkezinde, Coat. </li><li> Larva odasında mikroskop objektif bakan ventral yüzü koyun (nöromüsküler bileşke (NMJ) 26 ve 27 görüntülü sağlar) (Şekil 1 AE). </li><li> Grid bakmalıdır (Şekil 1 A) hava deliklerini, yağ tabakası üzerine plastik ara yerleştirin. Bu aşamada kontrol edin: spacer yüksekliği kabaca yarısı larva çapı, yarık genişliği larva çapının yaklaşık iki katı olmalıdır. </li><li> Mikroskop görüntüleme odasının yerleştirin </li></ol> B) Anesthetization larva <ol start="7"><li> Uygun bir anesthetization cihaz / buharlaştırıcı odasının iki girişler bağlayın. Up-front kontrol edin: Oda hava desfluran konsantrasyonu etkin güvenlik yönetmeliklerine belirtilen asla aşmamalıdır! Desfluran, sadece çok küçük bir toplam hacmi larvaları uyutmak için gerekli. 1 desfluran% 15 uygulaması (v / v) bir deneyde kullanılmak üzere ideal konsantrasyonu belirlemek için iyi bir başlangıç ​​noktası olarak kanıtlamıştır. Güvenlik nedenleri için buharlaştırıcı in vivo görüntüleme 2-4 saat için gerekli daha asla desfluran içermesi gerektiğini öneriyoruz. </li><li> Diğer ucunda bir bardak su odasının prizine takılı tüp daldırın sonra yaklaşık beş saniye süreyle odasına anestezi akışını kontrol vanalarının açık. Bu aşamada kontrol edin: Hava kabarcıkları su yükselirken musunuz? Değilse odasına sızdıran. </li><li> Yaklaşık üç saniye vanalarını kapatın. </li><li> Mikroskop kalıntı larva hareketleri izleyin. Kalp atışı ve kas hareketleri de kontrol edin. Larva anesthetization tamamlanana kadar gerekli açık ve yakın vanalar adım 8 ve 9 olarak açıklanan, daha sonra tüm vanaları kapatın ve görüntüleme başlayın. Bu aşamada kontrol edin: Artık kas hareketi ya da kalp atışı, larva düzgün anestezi olmadığını gösterir. Komple anesthetization yüksek çözünürlüklü görüntüler için hayati önem taşımaktadır. Larva normalde belirli bir zamanda daha uzun bir süre 15-20 dakika narkoz olmamalıdır. </li></ol> C) Görüntüleme <ol start="11"><li> Doğru pozisyonu ve görüntü faiz yapısı (Şekil 2-4) belirleyin. </li></ol> Den D) Kurtarma anesthetization <ol start="12"><li> Görüntüleme odasına hava izin tamamlandıktan sonra. Bu aşamada kontrol edin: Iletilen halojen ışık ile larva kalp atışı ve kas kasılmaları kontrol edin. Kasları sözleşme başlangıç ​​olarak görüntüleme odasından larva kaldırmak için güvenlidir. </li><li> Anesthetization cihazdan oda ayırın ve mikroskop çıkarın. Odanın dikkatlice sökün ve püresi sinek ekimi orta larva. </li><li> Bir kuluçka çanak uygun sıcaklıkta saklayın. </li></ol> E) Zaman serileri <ol start="15"><li> Yeterli zaman noktalarında elde edilmiştir kadar 2-14 adımları tekrarlayın. Alternatif protokolü: Larva, 30 dakikalık zaman aralığı içinde bir kereden fazla yansıması ise, bir sonraki anesthetization noktaya kadar görüntüleme odasında bırakmak için pratik olabilir. Yeterli zaman noktalarında elde edilmiştir kadar 7-14 adımları tekrarlayın. Bu aşamada kontrol edin: Larva, bir defada iki saatten fazla görüntüleme odasında tutulması, ne de bir seferde 15-20 dakika daha uzun süre narkoz için değildir. </li></ol><img src="/files/ftp_upload/2249/2249fig1.jpg" alt="Şekil 1" /> Görüntüleme odasının Şekil 1 Meclisi. (A) larva ve plastik ayırıcı yağ tabakası üzerine yerleştirin. (B) yerleştirin ve ayırıcı bir 22 x 22 mm kapak kayma (C) metal halka ile larva pozisyonunu düzeltmek ve (D) odasına yakın odasına pleksiglas kılavuz yüzük takmak. (E) Şimdi odanın mikroskop üzerine monte edilmek üzere hazırdır. <img src="/files/ftp_upload/2249/2249fig2.jpg" alt="Şekil 2" /> Şekil 2 Drosophila larvaları Vücut duvarı kasları. Kaslar ve NMJs CD8-GFP-Sh füzyon proteini 8 ifade görüntülendi. Larva haline manikür ventral tarafında odaklanarak zaman görüldüğü gibi, kaslar gösterilmektedir. (AC) en yüzeysel kas, 27, gösterilir, (KL) en iç kasları, 6 ve 7 gösterilir. Ölçek Bar: 100 mikron, dilimler arasında ΔZ 2 mikron. <img src="/files/ftp_upload/2249/2249fig3.jpg" alt="Şekil 3" /> Şekil 3 Drosophila larvaların vücut duvarı kasları Kimlik. L3 Drosophila larvaları, segment A3 kasları kimliği gösterilir. Kaslar ve NMJs CD8-GFP-Sh füzyon proteini 8 ifade görüntülendi. Larvaları manikür ventral tarafında odaklanarak zaman görüldüğü gibi, kaslar görüntülenir. En yüzeysel kas, 27, AC gösterilir KL en iç kasları, 6 ve 7 gösterilir. Ölçek Bar: 100 mikron, ΔZ dilimleri arasında 2 mikron. <img src="/files/ftp_upload/2249/2249fig4.jpg" alt="Şekil 4" /> Şekil 4 Kimlik Drosophila larvaların nöromüsküler kavşak. (AD) NMJs DGluRIIA-MRFP füzyon proteini 1 ifadesi ile görüntülendi. NMJs larva haline manikür ventral tarafında odaklanarak zaman gözlemlediği gibi gösterilmiştir. (AB) en yüzeysel NMJ, 27, gösterilen, en iç NMJs, 6 ve 7, CD gösterilmiştir. Ölçek Bar: 100 mikron, ΔZ dilimleri arasında 5 mikron. (E) ve (F) yüzeysel (E) ve (F) daha iç NMJs kimliği referans verilir.

<ol>
	<li>Rasse, T. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glutamate+receptor+dynamics+organizing+synapse+formation+in+vivo.">Glutamate receptor dynamics organizing synapse formation in vivo.</a> Nat Neurosci. 8, 898-905 (2005).</li><li>Rasse, T. 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> In vivo imaging of long-term changes in the Drosophila neuromuscular system [dissertation]. , (1988).</li><li>Fuger, P., Behrends, L. B., Mertel, S., Sigrist, S. J., Rasse, T. 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+synapse+development+and+measuring+protein+dynamics+using+two-color+fluorescence+recovery+after+photo-bleaching+at+Drosophila+synapses.">Live imaging of synapse development and measuring protein dynamics using two-color fluorescence recovery after photo-bleaching at Drosophila synapses.</a> Nat Protoc. 2, 3285-3298 (2007).</li><li>Wang, X., Schwarz, T. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+axonal+transport+of+mitochondria.">Imaging axonal transport of mitochondria.</a> Methods in enzymology. 457, 319-333 (2009).</li><li>Vinegoni, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mesoscopic+fluorescence+tomography+for+in-vivo+imaging+of+developing+Drosophila.">Mesoscopic fluorescence tomography for in-vivo imaging of developing Drosophila.</a> J Vis Exp. , (2009).</li><li>Lin, C. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Label-free+imaging+of+Drosophila+larva+by+multiphoton+autofluorescence+and+second+harmonic+generation+microscopy.">Label-free imaging of Drosophila larva by multiphoton autofluorescence and second harmonic generation microscopy.</a> Journal of biomedical optics. 13, 050502-050502 (2008).</li><li>Levitan, E. S., Lanni, F., Shakiryanova, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+imaging+of+vesicle+motion+and+release+at+the+Drosophila+neuromuscular+junction.">In vivo imaging of vesicle motion and release at the Drosophila neuromuscular junction.</a> Nat Protoc. 2, 1117-1125 (2007).</li><li>Zito, K., Parnas, D., Fetter, R. D., Isacoff, E. Y. &. a. m. p. ;. a. m. p., Goodman, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Watching+a+synapse+grow:+noninvasive+confocal+imaging+of+synaptic+growth+in+Drosophila.">Watching a synapse grow: noninvasive confocal imaging of synaptic growth in Drosophila.</a> Neuron. 22, 719-729 (1999).</li></ol>

Sunulan yöntem başlangıçta Drosophila melanogaster larvaların vücut duvarı kasları glutamaterjik sinapsların çalışma için geliştirilmiştir. Drosophila nöromüsküler bileşke (NMJ), kas ve nöronlar basmakalıp bir sito-mimarisi ile karakterizedir ve bu nedenle in vivo görüntüleme için ideal uygundur . Ancak, açıklanan anesthetization protokol görüntüleme NMJ sınırlı değil; Drosophila larvaların şeffaflık, organların gelişimini incelemek için açıklanan protokol adaptasyon, hücre göçü, hücre içinde aksonal kargo ve alt-hücresel yeniden yapılanma taşınması kolaylaştırır .

Reaktifler: <ul><li> Desfluran (Suprane, Baxter, Unterschleißheim, Almanya) </li><li> Halokarbon yağı (örneğin Voltalef H10S yağ / Atofina, Puteaux, Fransa) </li><li> Fly kültür ortamı </li></ul> Ekipman: <ul><li> Ters konfokal mikroskop </li><li> Binoküler mikroskop (örneğin STEMI 2000, Carl Zeiss Jena, Almanya) </li><li> Sıralamak olarak kullanılan küçük bir fırça uçar </li><li> Kuvöz </li><li> Özel inşa görüntüleme odası (mekanik çizimler için bakiniz Ref 3) </li><li> Buharlaştırıcı / anesthetization cihazı (Ayrıntılı bilgi için Referans 3) </li></ul>

in vivo imaging of intact drosophila larvae at sub-cellular resolution

Istenen transgenik hayvan hatlarının verimli kurulması sağlayan optik görüntüleme, genetik olarak kodlanmış fluorophores ve genetik araçları son gelişmeler bir yaşam bağlamında incelenecek biyolojik süreçlerin etkin ve hatta bazı durumlarda organizma davranıyor. Bu protokolde, alt hücresel çözünürlük 1-3 sinaptik popülasyonlarının geliştirme ve plastisite takip desfluran uçucu anestezik kullanarak sağlam Drosophila larvaları, uyutmak nasıl anlatacağız . Drosophila melanogaster larvaları uyutmak için diğer yararlı yöntemler, daha önce açıklanan 4,5,6,7,8 alınmış olsa da, burada sunulan protokol kombine aşağıdaki temel özellikleri nedeniyle önemli gelişmeler göstermektedir: (1) anesthetization çok yüksek derecede, hatta kalp atışı gün 2 (beşten fazla zaman puan saatlik bir süre içinde kayıt izin&gt; anesthetization siklus başına% 90 (2), yüksek hayatta kalma oranı, 150 nm ye kadar 1 lateral çözünürlük izin tutuklandı 3) 2 durumlarda bize sağlayan, yüksek hassasiyetli fizyolojik düzeyde ifade edilen proteinlerin dinamiklerini incelemek için. Ayrıntılı olarak GluR-IIA istikrarlı transgenik hatları endojen organizatörü 1 ve ekzon tuzak hattı FasII-GFP 1 ile ifade postsinaptik glutamat reseptör altbirim görselleştirmek için başardık. (4) 4,7 larvaları diğer yöntemlerin aksine sadece hayatta değil görüntülü, aynı zamanda sağlam olabilir (yani olmayan disseke) gözlem gün 1 bir dizi üzerinden gerçekleşmesini sağlar. Ekteki video, in vivo görüntüleme odasının 2,3 bireysel parçaların fonksiyonu, larvaları, anesthetization prosedürü nasıl bir larva içinde belirli konumlarda yeniden tanımlamak için ve larvalarının güvenli kaldırma doğru montaj görüntüleme odası.

Watch this Scientific Journal Video about In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution at JoVE.com

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

Bu protokol, bozulmamış anesthetization ve görüntüleme için güvenilir bir yöntem açıklanır<em&gt; Drosophila melanogaster</em&gt; Larvaları. Biz bir kaç gün öncesine kadar volatil anestetik desfluran yapıların alt hücresel çözünürlük ve yeniden tanımlama tekrarlayan görüntüleme için izin kullanmış olması<sup&gt; 1</sup&gt;.

Alt hücresel Çözünürlük bozulmamış Drosophila Larva in vivo Görüntüleme

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

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Biology

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

14.5K Görüntüleme.  University of Tübingen. Bu protokol, bozulmamış anesthetization ve görüntüleme için güvenilir bir yöntem açıklanır Drosophila melanogaster Larvaları. Biz bir kaç gün öncesine kadar volatil anestetik desfluran yapıların alt hücresel çözünürlük ve yeniden tanımlama tekrarlayan görüntüleme için izin kullanmış olması 1.

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

High-resolution Measurement of Odor-Driven Behavior in Drosophila Larvae

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral paradigm, the experimenter usually assumes that the rapid diffusion of odorant molecules from the source leads to the creation of a stable gradient in the dish. To establish a quantitative correlation between sensory inputs and behavioral responses, it is necessary to achieve a more thorough characterization of the odorant stimulus conditions. In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

Yüksek çözünürlüklü Ölçüm Drosophila Larva Koku-Driven Davranış

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral ...

Biyoloji

Labeling hESCs and hMSCs with Iron Oxide Nanoparticles for Non-Invasive in vivo Tracking with MR Imaging

In recent years, stem cell research has led to a better understanding of developmental biology, various diseases and its potential impact on regenerative medicine. A non-invasive method to monitor the transplanted stem cells repeatedly in vivo would greatly enhance our ability to understand the mechanisms that control stem cell death and identify trophic factors and signaling pathways that improve stem cell engraftment. MR imaging has been proven to be an effective tool for the in vivo depiction of stem cells with near microscopic anatomical resolution. In order to detect stem cells with MR, the cells have to be labeled with cell specific MR contrast agents. For this purpose, iron oxide nanoparticles, such as superparamagnetic iron oxide particles (SPIO), are applied, because of their high sensitivity for cell detection and their excellent biocompatibility. SPIO particles are composed of an iron oxide core and a dextran, carboxydextran or starch coat, and function by creating local field inhomogeneities, that cause a decreased signal on T2-weighted MR images. This presentation will demonstrate techniques for labeling of stem cells with clinically applicable MR contrast agents for subsequent non-invasive in vivo tracking of the labeled cells with MR imaging.

For the evaluation of new stem cell therapies it is important to non-invasively track the injected cells in vivo. This video will show you how to label human mesenchymal and embryonic stem cells with iron oxide based contrast agents in vivo for subsequent MR imaging in vivo.

Demir Oksit hESC ve hMSCs Etiketleme MR Görüntüleme in vivo Takip Non-Invasive Nanopartiküller

In recent years, stem cell research has led to a better understanding of developmental biology, various diseases and its potential impact on regenerative ...

High-Resolution Video Tracking of Locomotion in Adult Drosophila Melanogaster

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field. Studying locomotor behavior in Drosophila melanogaster relies on the ability to be able to quantify changes in motion during or in response to a given task. For this reason, a high-resolution video tracking system, such as the one we describe in this paper, is a valuable tool for measuring locomotion in real-time. Our protocol involves the use of an initial air pulse to break the flies momentum, followed by a thirty second filming period in a square chamber. A tracking program is then used to calculate the instantaneous speed of each fly within the chamber in 10 msec increments. Analysis software then compiles this data, and outputs a variety of parameters such as average speed, max speed, time spent in motion, acceleration, etc. This protocol will discuss proper feeding and management of flies for behavioral tasks, handling flies without anesthetization or immobilization, setting up a controlled environment, and running the assay from start to finish.

The study of complex locomotor behavior in Drosophila melanogaster is dependent upon the ability to quantify changes in a given fly's movement. This article demonstrates how to do this using a high-resolution tracking system.

Yetişkin Drosophila melanogaster Locomotion Yüksek Çözünürlüklü Video Takip

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field.  Studying ...

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Visualizing developing organ formation as well as progession and treatment of disease often heavily relies on the ability to optically interrogate molecular and functional changes in intact living organisms. Most existing optical imaging methods are inadequate for imaging at dimensions that lie between the penetration limits of modern optical microscopy (0.5-1mm) and the diffusion-imposed limits of optical macroscopy (&gt;1cm) [1]. Thus, many important model organisms, e.g. insects, animal embryos or small animal extremities, remain inaccessible for in-vivo optical imaging. 
Although there is increasing interest towards the development of nanometer-resolution optical imaging methods, there have not been many successful efforts in improving the imaging penetration depth. The ability to perform in-vivo imaging beyond microscopy limits is in fact met with the difficulties associated with photon scattering present in tissues. Recent efforts to image entire embryos for example [2,3] require special chemical treatment of the specimen, to clear them from scattering, a procedure that makes them suitable only for post-mortem imaging. These methods however evidence the need for imaging larger specimens than the ones usually allowed by two-photon or confocal microscopy, especially in developmental biology and in drug discovery.
We have developed a new optical imaging technique named Mesoscopic Fluorescence Tomography [4], which appropriate for non-invasive in-vivo imaging at dimensions of 1mm-5mm. The method exchanges resolution for penetration depth, but offers unprecedented tomographic imaging performance and it has been developed to add time as a new dimension in developmental biology observations (and possibly other areas of biological research) by imparting the ability to image the evolution of fluorescence-tagged responses over time. As such it can accelerate studies of morphological or functional dependencies on gene mutations or external stimuli, and can importantly, capture the complete picture of development or tissue function by allowing longitudinal time-lapse visualization of the same, developing organism.
The technique utilizes a modified laboratory microscope and multi-projection illumination to collect data at 360-degree projections. It applies the Fermi simplification to Fokker-Plank solution of the photon transport equation, combined with geometrical optics principles in order to build a realistic inversion scheme suitable for mesoscopic range. This allows in-vivo whole-body visualization of non-transparent three-dimensional structures in samples up to several millimeters in size.
We have demonstrated the in-vivo performance of the technique by imaging three-dimensional structures of developing Drosophila tissues in-vivo and by following the morphogenesis of the wings in the opaque Drosophila pupae in real time over six consecutive hours.

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Mesoscopic fluorescence tomography operates beyond the penetration limits of tissue-sectioning fluorescence microscopy. The technique is based on multi-projection illumination and a photon transport description. We demonstrate in-vivo whole-body 3D visualization of the morphogenesis of GFP-expressing wing imaginal discs in Drosophila melanogaster.

Mezoskopik Floresans Tomografi için In-vivo Görüntüleme Geliştirme Drosophila</em

Visualizing  developing organ formation as well as progession and treatment of disease often  heavily relies on the ability to optically interrogate ...

In-vivo Centrifugation of Drosophila Embryos

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in buoyant density. However, when cells are disrupted prior to centrifugation, proteins and organelles in this non-native environment often inappropriately stick to each other. Here we describe a method to separate organelles by density in intact, living Drosophila embryos. Early embryos before cellularization are harvested from population cages, and their outer egg shells are removed by treatment with 50% bleach. Embryos are then transferred to a small agar plate and inserted, posterior end first, into small vertical holes in the agar. The plates containing embedded embryos are centrifuged for 30 min at 3000g. The agar supports the embryos and keeps them in a defined orientation. Afterwards, the embryos are dug out of the agar with a blunt needle. Centrifugation separates major organelles into distinct layers, a stratification easily visible by bright-field microscopy. A number of fluorescent markers are available to confirm successful stratification in living embryos. Proteins associated with certain organelles will be enriched in a particular layer, demonstrating colocalization. Individual layers can be recovered for biochemical analysis or transplantation into donor eggs. This technique is applicable for organelle separation in other large cells, including the eggs and oocytes of diverse species.

In-vivo Centrifugation of Drosophila Embryos

We describe a method to separate organelles by density in living Drosophila embryos. Embryos are embedded in agar and centrifuged. This technique yields reproducible separation of major organelles along the anterior-posterior embryo axis. This method facilitates colocalization experiments and yields organelle fractions for biochemical analysis and transplantation experiments.

In-vivo Drosophila Embriyolar Santrifüj

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in ...

Imaging Cell Shape Change in Living Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging. Cell shape changes in the fly are analogous to those in higher organisms, and they drive tissue morphogenesis. So, in many cases, their study has direct implications for understanding human disease (Table 1)1-5. On the sub-cellular scale, these cell shape changes are the product of activities ranging from gene expression to signal transduction, cell polarity, cytoskeletal remodeling and membrane trafficking. Thus, the Drosophila embryo provides not only the context to evaluate cell shape changes as they relate to tissue morphogenesis, but also offers a completely physiological environment to study the sub-cellular activities that shape cells.
The protocol described here is designed to image a specific cell shape change called cellularization. Cellularization is a process of dramatic plasma membrane growth, and it ultimately converts the syncytial embryo into the cellular blastoderm. That is, at interphase of mitotic cycle 14, the plasma membrane simultaneously invaginates around each of ~6000 cortically anchored nuclei to generate a sheet of primary epithelial cells. Counter to previous suggestions, cellularization is not driven by Myosin-2 contractility6, but is instead fueled largely by exocytosis of membrane from internal stores7. Thus, cellularization is an excellent system for studying membrane trafficking during cell shape changes that require plasma membrane invagination or expansion, such as cytokinesis or transverse-tubule (T-tubule) morphogenesis in muscle.
Note that this protocol is easily applied to the imaging of other cell shape changes in the fly embryo, and only requires slight adaptations such as changing the stage of embryo collection, or using "embryo glue" to mount the embryo in a specific orientation (Table 1)8-19. In all cases, the workflow is basically the same (Figure 1). Standard methods for cloning and Drosophila transgenesis are used to prepare stable fly stocks that express a protein of interest, fused to Green Fluorescent Protein (GFP) or its variants, and these flies provide a renewable source of embryos. Alternatively, fluorescent proteins/probes are directly introduced into fly embryos via straightforward micro-injection techniques9-10. Then, depending on the developmental event and cell shape change to be imaged, embryos are collected and staged by morphology on a dissecting microscope, and finally positioned and mounted for time-lapse imaging on a confocal microscope.

Imaging Cell Shape Change in Living Drosophila Embryos

Early development of the fruit fly, Drosophila melanogaster, is characterized by a number of cell shape changes that are well suited for imaging approaches. This article will describe basic tools and methods required for live confocal imaging of Drosophila embryos, and will focus on a cell shape change called cellularization.

Living Görüntüleme Hücre Şekli Değiştir Drosophila Embriyolar

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging.  Cell shape ...

Microgavage of Zebrafish Larvae

The zebrafish has emerged as a powerful model organism for studying intestinal development1-5, physiology6-11, disease12-16, and host-microbe interactions17-25. Experimental approaches for studying intestinal biology often require the in vivo introduction of selected materials into the lumen of the intestine. In the larval zebrafish model, this is typically accomplished by immersing fish in a solution of the selected material, or by injection through the abdominal wall. Using the immersion method, it is difficult to accurately monitor or control the route or timing of material delivery to the intestine. For this reason, immersion exposure can cause unintended toxicity and other effects on extraintestinal tissues, limiting the potential range of material amounts that can be delivered into the intestine. Also, the amount of material ingested during immersion exposure can vary significantly between individual larvae26. Although these problems are not encountered during direct injection through the abdominal wall, proper injection is difficult and causes tissue damage which could influence experimental results. We introduce a method for microgavage of zebrafish larvae. The goal of this method is to provide a safe, effective, and consistent way to deliver material directly to the lumen of the anterior intestine in larval zebrafish with controlled timing. Microgavage utilizes standard embryo microinjection and stereomicroscopy equipment common to most laboratories that perform zebrafish research. Once fish are properly positioned in methylcellulose, gavage can be performed quickly at a rate of approximately 7-10 fish/ min, and post-gavage survival approaches 100% depending on the gavaged material. We also show that microgavage can permit loading of the intestinal lumen with high concentrations of materials that are lethal to fish when exposed by immersion. To demonstrate the utility of this method, we present a fluorescent dextran microgavage assay that can be used to quantify transit from the intestinal lumen to extraintestinal spaces. This test can be used to verify proper execution of the microgavage procedure, and also provides a novel zebrafish assay to examine intestinal epithelial barrier integrity under different experimental conditions (e.g. genetic manipulation, drug treatment, or exposure to environmental factors). Furthermore, we show how gavage can be used to evaluate intestinal motility by gavaging fluorescent microspheres and monitoring their subsequent transit. Microgavage can be applied to deliver diverse materials such as live microorganisms, secreted microbial factors/toxins, pharmacological agents, and physiological probes. With these capabilities, the larval zebrafish microgavage method has the potential to enhance a broad range of research fields using the zebrafish model system.

We present a novel method for microgavage of larval zebrafish utilizing standard embryo microinjection and stereomicroscopy equipment. We demonstrate that microgavage is a safe and efficient technique useful for delivering controlled amounts of diverse materials specifically into the larval zebrafish intestinal lumen.

Zebra balığı larvalarının Microgavage

The zebrafish has emerged as a powerful model organism for studying intestinal development1-5, physiology6-11, disease12-16, and host-microbe ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

Apoptotik hücresi Temizliği görüntülenmesi sırasında Canlı<em&gt; Drosophila</em&gt; Embriyo

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Ex vivo Method for High Resolution Imaging of Cilia Motility in Rodent Airway Epithelia

An ex vivo technique for imaging mouse airway epithelia for quantitative analysis of motile cilia function important for insight into mucociliary clearance function has been established. Freshly harvested mouse trachea is cut longitudinally through the trachealis muscle and mounted in a shallow walled chamber on a glass-bottomed dish. The trachea sample is positioned along its long axis to take advantage of the trachealis muscle to curl longitudinally. This allows imaging of ciliary motion in the profile view along the entire tracheal length. Videos at 200 frames/sec are obtained using differential interference contrast microscopy and a high speed digital camera to allow quantitative analysis of cilia beat frequency and ciliary waveform. With the addition of fluorescent beads during imaging, cilia generated fluid flow also can be determined. The protocol time spans approximately 30 min, with 5 min for chamber preparation, 5-10 min for sample mounting, and 10-15 min for videomicroscopy.

Ex vivo Method for High Resolution Imaging of Cilia Motility in Rodent Airway Epithelia

An easy and reliable technique for visualizing and quantifying airway cilia motility and cilia generated flow using mouse trachea is described. This technique can be modified to determine how a wide range of factors influence cilia motility, including pharmacological agents, genetic factors, environmental exposures, and/or mechanical factors such as mucus load.

Kemirgen Havayolu Epiteli içinde Cilia Motilite Yüksek Çözünürlüklü Görüntüleme için ex vivo Yöntemi

An ex vivo technique for imaging mouse airway epithelia for quantitative  analysis of motile cilia function important for insight into mucociliary  ...

Confocal Imaging of Single Mitochondrial Superoxide Flashes in Intact Heart or In Vivo

Mitochondrion is a critical intracellular organelle responsible for energy production and intracellular signaling in eukaryotic systems. Mitochondrial dysfunction often accompanies and contributes to human disease. Majority of the approaches that have been developed to evaluate mitochondrial function and dysfunction are based on in vitro or ex vivo measurements. Results from these experiments have limited ability in determining mitochondrial function in vivo. Here, we describe a novel approach that utilizes confocal scanning microscopy for the imaging of intact tissues in live aminals, which allows the evaluation of single mitochondrial function in a real-time manner in vivo. First, we generate transgenic mice expressing the mitochondrial targeted superoxide indicator, circularly permuted yellow fluorescent protein (mt-cpYFP). Anesthetized mt-cpYFP mouse is fixed on a custom-made stage adaptor and time-lapse images are taken from the exposed skeletal muscles of the hindlimb. The mouse is subsequently sacrificed and the heart is set up for Langendorff perfusion with physiological solutions at 37 &deg;C. The perfused heart is positioned in a special chamber on the confocal microscope stage and gentle pressure is applied to immobilize the heart and suppress heart beat induced motion artifact. Superoxide flashes are detected by real-time 2D confocal imaging at a frequency of one frame per second. The perfusion solution can be modified to contain different respiration substrates or other fluorescent indicators. The perfusion can also be adjusted to produce disease models such as ischemia and reperfusion. This technique is a unique approach for determining the function of single mitochondrion in intact tissues and in vivo.

Confocal Imaging of Single Mitochondrial Superoxide Flashes in Intact Heart or In Vivo

Confocal scanning microscopy is applied for imaging single mitochondrial events in perfused heart or skeletal muscles in live animal. Real-time monitoring of single mitochondrial processes such as superoxide flashes and membrane potential fluctuations enables the evaluation of mitochondrial function in a physiologically relevant context and during pathological perturbations.

Tek Mitokondriyal süperoksit Bozulmamış Kalp Flaşlar veya konfokal Görüntüleme<em&gt; In Vivo</em

Mitochondrion is a critical intracellular organelle responsible for energy production and intracellular signaling in eukaryotic systems. Mitochondrial ...