SG는 버팔로의 뉴욕 주립 대학에서와 존 R. Oishei 재단 기금에 의해 지원됩니다.

1. 시약의 준비 <ol><li> 128 MM NaCl, CaCl 4mm짜리 2, 4mm짜리 MgCl 2, 2 KCl MM, 5mM HEPES 및 36mM 자당를 사용하여 절개 1X 버퍼를 준비합니다. PH 7.2 솔루션 및 필터 소독. </li><li> 16 % 포름 알데히드 및​​ 1X 해부 버퍼의 1시 1분 희석을 사용하여 정착액을 준비합니다. </li><li> 1X 인산을 사용하여 1X PBT 준비 살린 (PBS)는 1:100 비율 7.2 그리고 Triton X - 100의 PH에 버퍼. </li><li> 준비 5% 소 혈청 0.01 %가 1X PBS에서 Azide 나와 함께 (BSA) 알부민. </li></ol> 2. 해부를위한 준비 <ol><li> Sylgard 요리는 절개에 사용됩니다. Sylgard 184 실리콘 엘라스토머 기지를 따르고하고 페트리 접시에 에이전트 혼합물을 치료하여 요리를 준비합니다. 혼합물이 완전히 사용하기 전에 응고 및 건조 수 있습니다. </li><li> 절개 전에 깨끗한 # 5 포셉의 쌍, 직선 블레이드 vannas 가위의 클린 쌍, 그리고 핀을를 수집합니다. </li></ol> 3. 제 3 Instar의 애벌레의 해부 <ol><li> 배양 접시에 특정 유전자형 3 차 instar의 애벌레를 헤매고 수집합니다. </li><li> 음식 없애 탈이온수에서 애벌레를 씻으십시오. </li><li> 등의 측면까지 함께 sylgard 요리에 유충 놓으십시오. 그 앞쪽에 및 사후 끝에 유충을 핀. </li><li> 고정된 유충에 1X 해부 버퍼의 드롭 놓습니다. 고급 가위 따다에게 사후 끝 근처 등의 정중선에 유충을 사용합니다. 가위를 사용하여 앞의 끝에 통해 사후 하순부터 애벌레 표피를 잘라. </li><li> 해부 유충에 새로운 1X 해부 버퍼의 드롭 놓습니다. PIN을 사용하여 유충의 중간 근처 표피 중 하나 측면 가장자리를 선택하고 표피를 핀. 다른 핀 사용으로 그림과 같이 표피의 두 번째 측면 가장자리를 핀. 두 핀이 각 측면 측면에 사용됩니다. 아래 다이어그램을 참조하십시오. </li><li> 조심스럽게 첨부 segmental 신경 함께 두 개의 광학 엽 (叶)과 복부 신경만을 두뇌를 떠나, 창자, 지방 단체 및 기관을 제거하십시오. 절차의 나머지 부분은 sylgard 요리에 이루어집니다. </li></ol> 4. 해부 유충의 고정 <ol><li> 매 15 분 수정을 대체하여, 30 분 수정의 해부 유충 부화. </li><li> 고정 후, 10 분 간격으로 버퍼를 교체하여 30 분 1X PBT의 유충을 씻어. 유충은 항상 버퍼에 있​​어야합니다하는 것이 중요합니다. </li></ol> 5. 해부 유충의 항체 스테 이닝 <ol><li> 1X PBT에서 나 Azide와 5 % BSA의 적절한 농도를 diluting하여 기본 항체를 준비합니다. 최적의 농도는 다른 항체를 위해 다를 수 있습니다. </li><li> 마지막 1X PBT은 준비된 차 항체 솔루션 린스, 4 ° C 하룻밤에 습기가 실내에 품어 바꿉니다. 습기 챔버가 물에 젖었 김 - 잎사귀와 플라스틱 접시를 안감으로 구성되어 있습니다. </li><li> 그 다음날, 10 분 간격으로 버퍼를 교체하여 30 분 1X PBT에서 해부하는 유충을 씻어. </li><li> 1X PBT에서 나 Azide와 5 % BSA의 적절한 농도를 diluting하여 이차 항체를 준비하고, 한 시간 동안 25 ° C에서 차 항체 솔루션 유충 부화. 이차 항체가 민감한 빛 있으므로, 빛을 방지하기 위해 호일에 습한 챔버를 바꿈. </li><li> 부화 후 10 분 간격으로 버퍼를 교체하여 30 분 1X PBT의 유충을 씻어. </li></ol> 6. 장착 및 해부 유충의 시각화 <ol><li> 슬라이드에있는 유충을 설치하기 전에 (그림 참조)에 앉을 수있는 커버 슬립을위한 충분한 공간, 중간에 매니큐어 두 수직선을 확산하여 슬라이드를 준비합니다. 완전히 손톱 폴란드 건조하자. </li><li> 슬라이드에서 세로 매니큐어 라인 (그림 참조) 사이의 공간에 장착 버퍼의 한 방울을 넣으십시오. 슬라이드 이제 설치 준비가되었습니다. <img src="/files/ftp_upload/2128/2128.6.2.1.jpg" alt="그림 1 단계 6.2" /> <img src="/files/ftp_upload/2128/2128.6.2.2.jpg" alt="그림 2 단계 6.2" /></li><li> 마지막 1X PBT는 린스 후, 부드럽게 측면 핀을 제거합니다. 표피 찢어하지 않도록주의한다. </li><li> 조심스럽게 두 남은 핀을 제거하고 집게로 유충을 선택하고 준비 슬라이드 수평 유충을 놓으십시오. 유충이 이상의 이성을 상실하고 지향 복부 측면가 없습니다 있는지 확인하십시오. </li><li> 부드럽게 위에 커버 용지를 놓고 (그림 참조) 매니큐어와 가장자리를 밀봉. 유충 지금은 형광 현미경으로 시각화를위한 준비가되었습니다. <img src="/files/ftp_upload/2128/2128.6.5.jpg" alt="그림 1 단계 6.5" /></li></ol> 7. 대표 결과 <img src="/files/ftp_upload/2128/2128fig1.jpg" alt="그림 1" /> 그림 1 : 야생 유형 larv에서 애벌레 segmental 신경의 대표 이미지(CSP, Zinsmaier 외. 1994) 시냅스 소포 단백질 시스테인 문자열 단백질에 대한 항체를 사용하여. segmental 신경이 원활하게 스테인드됩니다. 바 = 20μm <img src="/files/ftp_upload/2128/2128fig2.jpg" alt="그림 2" /> 그림 2 : 시냅스 소포 단백질 시스테인 문자열 단백질 (CSP)에 대한 항체를 사용하여 모터 단백질 돌연변이 유충에서 애벌레 segmental 신경의 대표 이미지. segmental 신경이 CSP (화살표)에 대한 밝은 얼룩 거대한 축적을 표시합니다.

<ol>
	<li>Gunawardena, S., Goldstein, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Disruption+of+Axonal+Transport+and+Neuronal+Viability+by+Amyoid+Precursor+Protein+Mutations+in+Drosophila.">Disruption of Axonal Transport and Neuronal Viability by Amyoid Precursor Protein Mutations in Drosophila.</a> Neuron. 32 (2), 389-401 (2001).</li><li>Gunawardena, S., Her, L. S., Brusch, R. G., Laymon, R. A., Niesman, I. R., Gordesky-Gold, B., Sintasath, L., Bonini, N. M., Goldstein, L. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Disruption+of+axonal+transport+by+loss+of+huntingtin+or+expression+of+pathogenic+polyQ+proteins+in+Drosophila.">Disruption of axonal transport by loss of huntingtin or expression of pathogenic polyQ proteins in Drosophila.</a> Neuron. 40 (1), 1-2 (2003).</li><li>Hurd, D. D., Saxton, W. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kinesin+mutations+cause+motor+neuron+disease+phenotypes+by+disrupting+fast+axonal+transport+in+Drosophila.">Kinesin mutations cause motor neuron disease phenotypes by disrupting fast axonal transport in Drosophila.</a> Genetics. 144 (3), 1075-1085 (1996).</li><li>Zinsmaier, K. E., Eberle, K. K., Buchner, E., Walter, N., Benzer, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Paralysis+and+early+death+in+cysteine+string+protein+mutants+of+Drosophila.">Paralysis and early death in cysteine string protein mutants of Drosophila.</a> Science. 263, 977-980 (1994).</li></ol>

관심 없음 충돌 선언하지 않습니다.

Drosophila 애벌레 segmental 신경 axonal 교통 메커니즘을 연구하기위한 강력한 시스템입니다. 3 차 instar 애벌레의 절개가 올바르게 수행하는 경우, 등 근육과 NMJs 같은 CNS, PNS, 및 관련 조직, 모든 구성 요소는 하나의 유충 이내에 검사를하실 수 있습니다. 우리는 허드와 색스턴 (1996)에 의해 출판 하나에서이 프로토콜을 적응있다. 이 프로토콜은 다른 항체의 연결을 테스트하는 적응 될 수 ...

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<td>Dumont #5 Forceps</td>
<td>&nbsp;</td>
<td>Fine Science Tools, Inc.</td>
<td>11252-20</td>
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<td>Minutien Pins, Stainless Steel</td>
<td>&nbsp;</td>
<td>Fine Science Tools, Inc.</td>
<td>26002-10</td>
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<td>Mcpherson-Vannas Straight Blade 8cm Scissors</td>
<td>&nbsp;</td>
<td>Fisher Scientific</td>
<td>50822236</td>
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<td>Vectashield, Mounting Medium</td>
<td>&nbsp;</td>
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<td>Alexa Fluor 568 Goat Anti-Mouse IgG</td>
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visualization of larval segmental nerves in 3rd instar drosophila larval preparations

Drosophila melanogaster 특히 때문에 편리 유전 완전 합성 게놈의 신경 계통의 발달과 기능을 연구를위한 강력한 모델 시스템으로 대두되고 있습니다. 또한, 애벌레 신경 시스템은 애벌레 segmental 신경이 신체의 길이를 따라 결말 두뇌 및 신경 터미널 내에있는 자신의 세포 기관과 axons의 번들을 포함하는 등 axonal 교통의 메커니즘을 연구하는 이상적인 모델 시스템입니다. 여기 애벌레 segmental 신경 사이 시냅스 소포 단백질의 시각화를위한 절차를 설명합니다. 제대로하면, 신경 시스템의 모든 구성 요소는 이러한 근육과 NMJs 등 관련 조직과 함께, 손상 손상 및 시각 준비가 남아있다. 다양한 변이를 갖고 제 3 instar의 애벌레가 해부되고, 고정 시냅스 소포 항체와 incubated, 시각 및 야생 입력 유충에 비해. 이 절차는 쉽게 애벌레 segmental 신경 내에서 볼 수 있습니다 단백질의 다양한 배포판에 여러 다른 시냅스의 연결 또는 항체 및 변경 적용할 수 있습니다.

Watch this Scientific Journal Video about Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations at JoVE.com

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

Drosophila melanogaster의 애벌레가 애벌레 segmental 신경 이내 axonal 수송의 메커니즘을 조사하기 위해 이상적인 모델 시스템을 제공합니다. 이 절차를 사용하여 다양한 변이를 갖고 제 3 instar의 애벌레는 야생 타입 애벌레 비교할 수 있습니다.

3 애벌레 Segmental 신경의 시각화 차 Instar Drosophila 애벌레 준비

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of ...

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Neuroscience

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

10.2K 조회수.  SUNY-University at Buffalo. Drosophila melanogaster의 애벌레가 애벌레 segmental 신경 이내 axonal 수송의 메커니즘을 조사하기 위해 이상적인 모델 시스템을 제공합니다. 이 절차를 사용하여 다양한 변이를 갖고 제 3 instar의 애벌레는 야생 타입 애벌레 비교할 수 있습니다.

비디오: Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different neurological diseases. Defects in this essential process can have detrimental effects on neuronal functioning and development. We have developed a dissection protocol that is designed to expose the Drosophila larval segmental nerves to view axonal transport in real time. We have adapted this protocol for live imaging from the one published by Hurd and Saxton (1996) used for immunolocalizatin of larval segmental nerves. Careful dissection and proper buffer conditions are critical for maximizing the lifespan of the dissected larvae. When properly done, dissected larvae have shown robust vesicle transport for 2-3 hours under physiological conditions. We use the UAS-GAL4 method 1 to express GFP-tagged APP or synaptotagmin vesicles within a single axon or many axons in larval segmental nerves by using different neuronal GAL4 drivers. Other fluorescently tagged markers, for example mitochrondria (MitoTracker) or lysosomes (LysoTracker), can be also applied to the larvae before viewing. GFP-vesicle movement and particle movement can be viewed simultaneously using separate wavelengths.

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

This protocol discusses the live dissection of Drosophila larvae for the purpose of imaging the movement of GFP tagged axonal vesicles on microtubule tracks.

Drosophila 애벌레 Segmental Axons 이내 시냅틱 Vesicles의 생체내 시각화에

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different ...

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신경과학

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

The Gal4/ UAS binary method is powerful for gene and neural circuitry manipulation in Drosophila. For most neurobiological studies, however, Gal4 expression is rarely tissue-specific enough to allow for precise correlation of the circuit with behavioral readouts. To overcome this major hurdle, we recently developed the FINGR method to achieve a more restrictive Gal4 expression in the tissue of interest. The FINGR method has three components: 1) the traditional Gal4/UAS system; 2) a set of FLP/FRT-mediated Gal80 converting tools; and 3) enhancer-trap FLP (ET-FLP). Gal4 is used to define the primary neural circuitry of interest. Paring the Gal4 with a UAS-effector, such as UAS-MJD78Q or UAS-Shits, regulates the neuronal activity, which is in turn manifested by alterations in the fly behavior. With an additional UAS-reporter such as UAS-GFP, the neural circuit involved in the specific behavior can be simultaneously mapped for morphological analysis. For Gal4 lines with broad expression, Gal4 expression can be restricted by using two complementary Gal80-converting tools: tubP&gt;Gal80&gt; ('flip out') and tubP&gt;stop&gt;Gal80 ('flip in'). Finally, investigators can turn Gal80 on or off, respectively, with the help of tissue-specific ET-FLP. In the flip-in mode, Gal80 will repress Gal4 expression wherever Gal4 and ET-FLP intersect. In the flip-out mode, Gal80 will relieve Gal4 repression in cells in which Gal4 and FLP overlap. Both approaches enable the restriction of the number of cells in the Gal4-defined circuitry, but in an inverse pattern. The FINGR method is compatible with the vast collection of Gal4 lines in the fly community and highly versatile for traditional clonal analysis and for neural circuit mapping. In this protocol, we demonstrate the mapping of FLP expression patterns in select ET-FLPx2 lines and the effectiveness of the FINGR method in photoreceptor cells. The principle of the FINGR method should also be applicable to other genetic model organisms in which Gal4/UAS, Gal80, and FLP/FRT are used.

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

We describe a Flippase-induced intersectional Gal80/Gal4 repression (FINGR) method, allowing tissue-specific FLP to determine Gal80 expression patterns. Wherever Gal4 and FLP overlap, Gal4 expression is turned on (Gal80 flipped out) or off (Gal80 flipped in). The FINGR method is versatile for clonal analysis and neural circuit mapping.

매핑 및 애벌레와 성인의 확장기 - 트랩 Flippase의 표현의 응용 Drosophila CNS

The Gal4/ UAS binary method is powerful for gene and neural circuitry manipulation in Drosophila. For most neurobiological studies, however, Gal4 ...

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including lens-secreting cone cells, pigment cells, a bristle cell and eight photoreceptors (PRs) R1-R8 2. The PRs have specialized microvillar structures, the rhabdomeres, which contain light-sensitive pigments, the Rhodopsins (Rhs). The rhabdomeres of six PRs (R1-R6) form a trapezoid and contain Rh1 3 4. The rhabdomeres of R7 and R8 are positioned in tandem in the center of the trapezoid and share the same path of light. R7 and R8 PRs stochastically express different combinations of Rhs in two main subtypes5: In the 'p' subtype, Rh3 in pR7s is coupled with Rh5 in pR8s, whereas in the 'y' subtype, Rh4 in yR7s is associated with Rh6 in yR8s 6 7 8.
Early specification of PRs and development of ommatidia begins in the larval eye-antennal imaginal disc, a monolayer of epithelial cells. A wave of differentiation sweeps across the disc9 and initiates the assembly of undifferentiated cells into ommatidia10-11. The 'founder cell' R8 is specified first and recruits R1-6 and then R7 12-14. Subsequently, during pupal development, PR differentiation leads to extensive morphological changes 15, including rhabdomere formation, synaptogenesis and eventually rh expression.
In this protocol, we describe methods for retinal dissections and immunohistochemistry at three defined periods of retina development, which can be applied to address a variety of questions concerning retinal formation and developmental pathways. Here, we use these methods to visualize the stepwise PR differentiation at the single-cell level in whole mount larval, midpupal and adult retinas (Figure 1).

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

The Drosophila retina is a crystal-like lattice composed of a small number of cell types that are generated in a stereotyped manner 1. Its amenability to sophisticated genetic analysis allows the study of complex developmental programs. This protocol describes dissections and immunohistochemistry of retinas at three discrete developmental stages, with a focus on photoreceptor differentiation.

애벌레의, Pupal 및 성인의 해부 및 Immunohistochemistry<em&gt; Drosophila</em&gt; 망막

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including ...

Forebrain Electrophysiological Recording in Larval Zebrafish

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked seizures, epilepsy can be acquired as a result of an insult to the brain or a genetic mutation. Efforts to model seizures in animals have primarily utilized acquired insults (convulsant drugs, stimulation or brain injury) and genetic manipulations (antisense knockdown, homologous recombination or transgenesis) in rodents. Zebrafish are a vertebrate model system1-3 that could provide a valuable alternative to rodent-based epilepsy research. Zebrafish are used extensively in the study of vertebrate genetics or development, exhibit a high degree of genetic similarity to mammals and express homologs for ~85% of known human single-gene epilepsy mutations. Because of their small size (4-6 mm in length), zebrafish larvae can be maintained in fluid volumes as low as 100 &mu;l during early development and arrayed in multi-well plates. Reagents can be added directly to the solution in which embryos develop, simplifying drug administration and enabling rapid in vivo screening of test compounds4. Synthetic oligonucleotides (morpholinos), mutagenesis, zinc finger nuclease and transgenic approaches can be used to rapidly generate gene knockdown or mutation in zebrafish5-7. These properties afford zebrafish studies an unprecedented statistical power analysis advantage over rodents in the study of neurological disorders such as epilepsy. Because the "gold standard" for epilepsy research is to monitor and analyze the abnormal electrical discharges that originate in a central brain structure (i.e., seizures), a method to efficiently record brain activity in larval zebrafish is described here. This method is an adaptation of conventional extracellular recording techniques and allows for stable long-term monitoring of brain activity in intact zebrafish larvae. Sample recordings are shown for acute seizures induced by bath application of convulsant drugs and spontaneous seizures recorded in a genetically modified fish.

A simple method to record extracellular field potentials in the larval zebrafish forebrain is described. The method provides a robust in vivo read-out of seizure-like activity. This technique can be used with genetically modified zebrafish larvae carrying epilepsy-related genes or seizures evoked by administration of convulsant drugs.

애벌레의 Zebrafish의 Forebrain Electrophysiological 기록

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked ...

Functional Analysis of the Larval Feeding Circuit in Drosophila

The serotonergic feeding circuit in Drosophila melanogaster larvae can be used to investigate neuronal substrates of critical importance during the development of the circuit. Using the functional output of the circuit, feeding, changes in the neuronal architecture of the stomatogastric system can be visualized. Feeding behavior can be recorded by observing the rate of retraction of the mouth hooks, which receive innervation from the brain. Locomotor behavior is used as a physiological control for feeding, since larvae use their mouth hooks to traverse across an agar substrate. Changes in feeding behavior can be correlated with the axonal architecture of the neurites innervating the gut. Using immunohistochemistry it is possible to visualize and quantitate these changes. Improper handling of the larvae during behavior paradigms can alter data as they are very sensitive to manipulations. Proper imaging of the neurite architecture innervating the gut is critical for precise quantitation of number and size of varicosities as well as the extent of branch nodes. Analysis of most circuits allow only for visualization of neurite architecture or behavioral effects; however, this model allows one to correlate the functional output of the circuit with the impairments in neuronal architecture.

Functional Analysis of the Larval Feeding Circuit in Drosophila

The feeding circuit in Drosophila melanogaster larvae serves a simple yet powerful model that allows changes in feeding rate to be correlated with alterations in the stomatogastric neural circuitry. This circuit is composed of central serotonergic neurons that send projections to the mouth hooks as well as the foregut.

의 애벌레 먹이 회로의 기능적 분석 초파리</i

The serotonergic  feeding circuit in Drosophila  melanogaster larvae can be used to investigate neuronal substrates of  critical importance during the ...

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

의 라이브 영상<em&gt; 초파리</em&gt; 애벌레 Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

Why Quantification Matters: Characterization of Phenotypes at the Drosophila Larval Neuromuscular Junction

Most studies on morphogenesis rely on qualitative descriptions of how anatomical traits are affected by the disruption of specific genes and genetic pathways. Quantitative descriptions are rarely performed, although genetic manipulations produce a range of phenotypic effects and variations are observed even among individuals within control groups.&#160;Emerging evidence shows that morphology, size and location of organelles play a previously underappreciated, yet fundamental role in cell function and survival. Here we provide step-by-step instructions for performing quantitative analyses of phenotypes at the Drosophila larval neuromuscular junction (NMJ). We use several reliable immuno-histochemical markers combined with bio-imaging techniques and morphometric analyses to examine the effects of genetic mutations on specific cellular processes. In particular, we focus on the quantitative analysis of phenotypes affecting morphology, size and position of nuclei within the striated muscles of Drosophila larvae. The Drosophila larval NMJ is a valuable experimental model to investigate the molecular mechanisms underlying the structure and the function of the neuromuscular system, both in health and disease. However, the methodologies we describe here can be extended to other systems as well.

Why Quantification Matters: Characterization of Phenotypes at the Drosophila Larval Neuromuscular Junction

Morphology, size and location of intracellular organelles are evolutionarily conserved and appear to directly affect their function. Understanding the molecular mechanisms underlying these processes has become an important goal of modern biology. Here we show how these studies can be facilitated by the application of quantitative techniques.

상기 표현형의 특성을 : 정량이 중요한 이유<em&gt; 초파리</em&gt; 애벌레 신경 근육 학 분기점

Most studies on morphogenesis rely on qualitative descriptions of how anatomical traits are affected by the disruption of specific genes and genetic ...

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat open surfaces is complex,&#160;including: pausing, turning, and meandering. This complexity in the repertoire of movement hinders detailed analysis of the events occurring during a single crawl stride cycle. To overcome this obstacle, linear agarose channels were made that constrain larval behavior to straight, sustained, rhythmic crawling. In principle, because agarose channels and the Drosophila larval body are both optically clear, the movement of larval structures labeled by genetically-encoded fluorescent probes can be monitored in intact, freely-moving larvae. In the past, larvae were placed in linear channels and crawling at the level of whole organism, segment, and muscle were analyzed1. In the future, larvae crawling in channels can be used for calcium imaging to monitor neuronal activity. Moreover, these methods can be used with larvae of any genotype and with any researcher-designed channel. Thus the protocol presented below is widely applicable for studies using the Drosophila larva as a model to understand motor control.

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

The Drosophila larva is a powerful model system to study neural control of behavior. This publication describes the use of linear agarose channels to elicit sustained bouts of linear crawling and methods to quantify the dynamics of larval structures during repetitive crawling behavior.

선형 아가로 오스 채널을 사용하여 공부하기<em&gt; 초파리</em&gt; 애벌레 크롤링 동작

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat ...

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

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

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

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

단일 셀 해상도 형광 애벌레 뇌 문화에 살고 있는 초파리 Circadian Clock의 이미징

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

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

The ability of insects to navigate toward odor sources is based on the activities of their first-order olfactory receptor neurons (ORNs). While a considerable amount of information has been generated regarding ORN responses to odorants, the role of specific ORNs in driving behavioral responses remains poorly understood. Complications in behavior analyses arise due to different volatilities of odorants that activate individual ORNs, multiple ORNs activated by single odorants, and the difficulty in replicating naturally observed temporal variations in olfactory stimuli using conventional odor-delivery methods in the laboratory. Here, we describe a protocol that analyzes Drosophila larval behavior in response to simultaneous optogenetic stimulation of its ORNs. The optogenetic technology used here allows for specificity of ORN activation and precise control of temporal patterns of ORN activation. Corresponding larval movement is tracked, digitally recorded, and analyzed using custom written software. By replacing odor stimuli with light stimuli, this method allows for a more precise control of individual ORN activation in order to study its impact on larval behavior. Our method could be further extended to study the impact of second-order projection neurons (PNs) as well as local neurons (LNs) on larval behavior. This method will thus enable a comprehensive dissection of olfactory circuit function and complement studies on how olfactory neuron activities translate in to behavior responses.

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

This protocol analyzes navigational behavior of Drosophila larva in response to simultaneous optogenetic stimulation of its olfactory neurons. Light of 630 nm wavelength is used to activate individual olfactory neurons expressing a red-shifted channel rhodopsin. Larval movement is simultaneously tracked, digitally recorded, and analyzed using custom-written software.