킬로바이트는 NIH 부여 GM54544에 의해 지원됩니다.

1 부 : 장비 및 소모품 <ol><li> 좋은 해부 현미경은 배아 해부가 필요합니다, 40X 배율은 maximally 배율을 증가 25X 눈길을 조각으로 제안합니다. </li><li> 좋은 포셉 (숫자 5) 배아와 태아의 devitellinization의 수동 선택이 필요합니다. </li><li> 미세 유리 바늘을 만들고 수정할 수있는 장비가 필요합니다. 아이고 - 유리 바늘은 절개가 필요합니다. 우리는 해부 (오래)에 대한 단단한 유리를 선호하지만, 표준 두꺼운 벽의 유리 튜브 (외경 1-1.5 mm)가 잘 작동합니다. 어떤 사람들은 10 + 앰프 autotransformer 배터리를 사용하여 1M NaOH의 목욕탕에서 원하는 선명도에 electrolyzed 미세 텅스텐 바늘을 사용합니다. 날카롭게 텅스텐 절개의 바늘들은 비교적 탄력과 오래 지속되는 장점이 있습니다. 중공 유리 바늘은 (클램프 전극 패치와 유사한 구성) 플라스틱 튜브에 연결된과 호수의 흡입 및 퇴학에 대한 해부뿐만 아니라 접착제의 통제 배달하는 동안 사용됩니다. 유리 pullers의 다양한 유리 바늘 제조에 사용할 수 있습니다. 컴퓨터 프로그램 pullers는 모양과 크기의 다양한 잡아 미리 수 있습니다. 브라운 - 플라밍 모델 것은 (서터 악기 주식 회사) 널리 선호하고 있습니다. 복합 현미경 (20 - 40X 목표)을 사용하기 전에 점검하고 전극을 수정할 수 있어야합니다. </li><li> 솔루션의 두 가지 유형은 일반적으로 목욕 해부 동안 배아 조직을 위해 사용됩니다 : 1) &quot;표준&quot;(1 월 및 월, 1976a) 또는 &quot;수정된 표준&quot;(Broadie, 2000) 살린스, 일반적으로 다른 무척추 동물 시스템에서 레코딩에 사용되는 솔루션을 기반으로 , 2) &quot;haemolymph처럼&quot;(HL) 살린스 (스튜어트 외 1994), 표준 호수와 Drosophila의 haemolymph에서 측정한 이온 농도 사이의 타협. 그것은 이러한 살린스 중 아무도 정확하게 생체내에 haemolymph의 화학 성분 (Broadie, 2000)을 모방하지 않습니다 언급해야합니다. 표준 생리가 포함되어 있습니다 (MM)을 : 135 NaCl, 5 KCl, 4 MgCl 2, 1.8 CaCl 2, 72 자당, 5 TES, 산도 7.2. 경험은 또한 electrophysiological 녹음에 적합하지만, 노출 조직의 건강을 보호하기 위해 가장 가능성이 상업적으로 준비 곤충 생리 (예 : 슈나이더의 곤충 미디어)을 고려할 수 있습니다. </li></ol> 2 부 : 엠브료 스테이징 및 해부 <ol><li> 최적의 달걀 수집, 젊은 (&lt;7 일), 건강한 남성 후 2-3 일 동안 신선한 누워있는 냄비에 여성 파리 (20-40) 유지합니다. 부설 냄비는 일반적으로 60mm의 한천 플레이트를 다루는 적절한 공기 순환을 제공하기 위해 천공 100 ML 플라스틱 비커 (트라이 붓고)로 이루어져 있습니다. 한천 플레이트 애플이나 한천과 함께 강화된 포도 주스가 포함되어 있습니다. 몇 가지 요리법이 존재하지만, 다음과 같은 일례는 다음과 같습니다 물 700 ML, 25~30g의 한천, 주스 집중 (포도 혹은 사과), 0.5g 메틸 paraben (P - hydroxymethylbenzoate) 300 ML, 그리고 30g의 설탕을. 물과 한천을 압력솥, 별도로 설탕, 주스와 P - hydroxymethylbenzoate을 끓여. 한천과 주스 솔루션을 믹스하고 신속하게 거품을 제거, 60mm 접시에 부어. </li><li> 이전 계란 컬렉션, 파리는 적어도 이틀 동안 적어도 하루에 두 번 변경 신선한 접시에 효모 붙여넣기 (4 ° C에 저장된 물을 9 ML에 7g 베이커의 효모)와 함께 공급됩니다. 일 3, 좋은 냄비는 시간당 100-200 계란을 생산해야합니다. 더 나은 계란 누워은 긁힌 접시의 한천과의 측면에 유지 냄비에 관찰됩니다. 많은 배아가에 누워 또는 흠집 옆에 것입니다. </li><li> 배아 많은 수의를 수집, 부드럽게 페인트를 사용하여 바구니에 한천 플레이트에서 배아를 전송합니다. 바구니는 15 또는 50 ML의 원심 분리기 튜브로 만들 수 있습니다. 뚜껑과 튜브의 상단 사이에 화면 (70 μm의 메쉬)를 함정에 충분한 면적을두고 아래를 잘라. DH 2 0으로 배아를 씻어 외부 chorion를 제거하는 신선한 50 % 백퍼센트 표백제의 페트리 접시에 넣어. 또는, 계란은 잘 집게를 사용하여 개별적으로 수집하고, 표백제의 드롭에 수동으로 배치하여 dechorionated 수 있습니다. 데 - chorionation 30 초에서 2 분 (신선한 표백제가 훨씬 빨라집니다)에 걸릴 수 있고, 해부 현미경 아래에서 모니터링해야합니다. chorion의 제거는 반짝 이는 투명 vitelline 막을 제공합니다. 표백 빠르게 devitellinized 조직을 파괴 등 Dechorionated 계란은 간략하게, 씻어서, 또는 호수에 배치해야합니다. </li><li> 그것은 신중하게 원하는 발달 시대에 배아 단계에 중요합니다. 정의된 Drosophila 배아의 단계 (1-17, 캄포스 - 오르테가와 Hartenstein, 1985) 모든 기능 신경근육학 개발이 늦은 단계 16 단계 17 발생으로 유용하지 않습니다. 따라서 배아가 25 개발 시간에서 개최 아르 ° C를 humidified 배양기에서 수정 이후에 시간 더 일반적으로, 달걀 누워 (AEL) 이후. 이러한 조건에서 embryogenesis 21 정도 + / - 25 ° C. 1 시간 </li><li> 초과 달걀하다 (계란에서 1-2 시간적절하게 세 아르들)은 다음 형태학의 기준 볼 수 있습니다. DH 2 O의 광범위한 세척 후 dechorionated 계란이 볼 수있는 플라스틱 문화 요리 (DH 2 O 미만)에 배치됩니다. 정확한 스테이징은 해부 현미경 (캄포스 - 오르테가와 Hartenstein, 1985)로 반사광을 사용하여 형태학의 기준으로 확인하실 수 있습니다. 부화 직전의 성숙 말기 태아 (22 - 24 AEL)은 일반적으로 표피와 부풀기도의 세분화에 의해 인정됩니다. 배아는 GFP balancers하고 적절한 필터와 형광 해부 현미경을 사용하여 genotyped 수 있습니다. </li><li> 해부 들어, dechorionated 및 발달 개최 배아는 호수의 드롭 (단계 1.5 참조) 아래 coverslip에 (최대 지느러미 쪽) 첨부합니다. 배아는 유리 micropipette 또는 텅스텐 바늘 앞쪽에 또는 사후 끝 부분에서 막을 펑쳐링 다음 부드럽게 멤브레인에서 배아를 자유롭게하여 vitelline 막에서 제거됩니다. 해부 준비에서 배아는 등의 표면 접속 (절개는 복부 신경 시스템과 실험 neuromusculature를 노출합니다 최대 도설 면을 coverslip에 위치해야합니다. 단, 배 (胚)가 분명 다른 접근을 촉진하기위한 다른 방법으로 위치 수 티슈.) </li><li> 초기 단계의 배아 (&lt;16 시간 AEL)은 유리 또는 polylysine (; Broadie 및 베이트, 1993a, B 폴리 - L - 라이신의 브롬화 수 소산​​)와 코팅 유리를 청소에 직접 연결합니다. 표피 형성 이전 다음 배아는 (&gt; 16 시간 AEL), 이상적으로 Sylgard - 코팅 (다우 코닝) coverslips에게 붙어 있어야합니다. 수상 polymerizing cyanoacrylate 수술이나 동물 접착제는 (Broadie와 베이트, 1993c, D)이 사용됩니다. 접착제는 입을 압력에 의해 제어되는 접착제의 흐름과 고무 튜브에 연결된 작은 유리 전극 피펫 (10-20 마이크로 미터 내경 팁)를 통해 전달됩니다. 접착제가 호수에 연락시 신속하게 polymerizes으로 케어는 접착제 배달하는 동안 이동해야합니다. 접착제가 이가의 이온은 중합 있어야으로 gluing가, 아니하거나 낮은 이가의 이온 농도와 호수에서 수행할 수 없습니다 또한 있습니다. 접착제 소량 먼저 단단히 머리와 꼬리를 첨부하는 데 사용됩니다. </li><li> 배아의 끝부분이 coverslip에 붙어되면, 절개는 유리 전극 또는 날카롭게 텅스텐 바늘 등의 정중선을 따라 이루어집니다. 이것은 부드럽게 최종 상처를 만들기 전에 절개의 선을 perforating로 쉽게 이루어집니다. 절개의 측면은 다음 부드럽게 배아 평면을 확산하기 위해 더 많은 접착제로 coverslip에 붙어 있습니다. 직감, 지방 기관과, 선택, 기관은 다음 고무 튜브 (Broadie 및 베이트 1993a - C)에 부착된 유리 피펫을 사용하여 흡입하여 제거를 포함한 내부 장기. 배아 이제 복부 중추 신경계와 표피 아래, 주변 신경, coverslip에 평평 첨부하고, 체세포 근육이 실험에 대해 노출되어야합니다. </li></ol>

배아의 정확한 준비로 인해 단지 몇 시간의 시간 코스 이상의 기능 속성의 급속한 성숙에 중요합니다. 몇 가지 문제는이 준비를 복잡하게. 첫째, 대부분의 연구자들은 초과 계란 단계 배아에 들이죠를 사용하지만, 계란 누워있는 시간이 서로 다른 조건 (Broadie 외. 1992)의 동물에서 동물로 매우 다를 수 있습니다. 특히, 제한된 다이어트 여성 전에 누워로 연장 기간 동안 수정된 달걀을 유지하는 경?...

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<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Embryo Collection Cages</td>
<td>&nbsp;</td>
<td>Genesee Scientific (www.flystuff.com)</td>
<td>59-100 (for 60 mm dish; other sizes available)</td>
<td>Cages can also be home made using punctured tri-pour beakers, as shown in video</td>
</tr>
<tr>
<td>Sylgard 184</td>
<td>&nbsp;</td>
<td>Dow Corning</td>
<td>Available from various companies</td>
<td>Surgical glue adheres better to sylgard-coated coverslips</td>
</tr>
<tr>
<td>Fine glass tubing, outer diameter 1.0-1.5 mm</td>
<td>&nbsp;</td>
<td>various</td>
<td>&nbsp;</td>
<td>For pulling into fine glass needles for dissection and tubes for glue delivery</td>
</tr>
<tr>
<td>Plastic tubing, to attach to glass pipette for mouth suction and glue application</td>
<td>&nbsp;</td>
<td>Tygon</td>
<td>&nbsp;</td>
<td>Tubing inner diameter needs to match glass outer diameter.</td>
</tr>		</tbody>
		</table>
		</div>

harvesting and preparing drosophila embryos for electrophysiological recording and other procedures

Drosophila는 배아 발달과 기능 모두의 신경 과학 연구를위한 최고의 유전자 모델입니다. 전통적으로,이 필드는 주로 독립적인 역사 및 과학 사회와 서로 꽤을 분리하고 있습니다. 그러나 이러한 일반적으로 서로 다른 분야 사이의 인터페이스는 기능 전기적 신호 특성과 신경 회로 형성의 마지막 단계에서 기능성 화학 시냅스의 분화 취득을 기본 발달 프로그램입니다. 이 인터페이스는 조사를위한 중요한 지역입니다. Drosophila에서 기능의 개발이 단계는 embryogenesis의 마지막 세번째 동안 &lt;8시간 기간 (25 ° C에서)하는 동안 발생합니다. 이렇게 늦게 발달 기간 동안 힘든, 스며들지 표피 표피의 증착에 수사 때문에하는 어려운 여겨졌다. 획기적인 사전은 로컬 말기 태아의 제어 절개를 사용하는 표피에 적용할 수있는 물 polymerizing 수술 접착제의 적용되었다. 지느러미 세로 절개로 배아가 실험 조사하기 위해 복부 신경 코드 및 신체 벽면 근육을 노출, 평면 마련하실 수 있습니다. 이 제도는 크게 배아 버렸네 형성에 악영향을하고, 따라서 버렸네 연결과 기능 시냅스 신호 속성의 사양과 차별을 규제하는 분자 메커니즘을 밝혀 유전자 돌연변이를 분리하고 특성화하는 데 사용되었습니다.

Watch this Scientific Journal Video about Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures at JoVE.com

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

이 기술은 immunohistochemistry 또는 electrophysiological 레코딩을위한 Drosophila 배아의 neuromusculature를 제공​​합니다. 그것은 신경근육학 개발 초기에 이벤트를 공부하거나 부화 수없는 돌연변이의 전기 생리학을 수행하는 데 유용합니다.

수확 및 준비 Drosophila</emElectrophysiological 녹화 및 기타 수속을위한> 태아

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite ...

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Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

13.3K 조회수.  University of Illinois. 이 기술은 immunohistochemistry 또는 electrophysiological 레코딩을위한 Drosophila 배아의 neuromusculature를 제공​​합니다. 그것은 신경근육학 개발 초기에 이벤트를 공부하거나 부화 수없는 돌연변이의 전기 생리학을 수행하는 데 유용합니다.

비디오: Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

Preparation of Neuronal Cultures from Midgastrula Stage Drosophila Embryos

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos and their dechorionation using bleach are demonstrated. Using a glass pipet attached to a mouth suction tube, we illustrate the removal of all cells from single embryos. The method for dispersing cells from each embyro into a small (5 l) drop of medium on an uncoated glass coverslip is demonstrated. A view through the microscope at 1 hour after plating illustrates the preferred cell density. Most of the cells that survive when grown in defined medium are neuroblasts that divide one or more times in culture before extending neuritic processes by 12-24 hours. A view through the microscope illustrates the level of neurite outgrowth and branching expected in a healthy culture at 2 days in vitro. The cultures are grown in a simple bicarbonate based defined medium, in a 5% CO2 incubator at 22-24&deg;C. Neuritic processes continue to elaborate over the first week in culture and when they make contact with neurites from neighboring cells they often form functional synaptic connections. Neurons in these cultures express voltage-gated sodium, calcium, and potassium channels and are electrically excitable. This culture system is useful for studying molecular genetic and environmental factors that regulate neuronal differentiation, excitability, and synapse formation/function.

This video demonstrates the preparation of primary neuronal cultures from midgastrula stage Drosophila embryos. Views of live cultures show cells 1 hour after plating and differentiated neurons after 2 days of growth in a bicarbonate-based defined medium. The neurons are electrically excitable and form synaptic connections.

Midgastrula 스테이지 Drosophila의 태아에서 문화의 연결 준비

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos ...

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생물학

Electrophysiological Methods for Recording Synaptic Potentials from the NMJ of Drosophila Larvae

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. The larval neuromuscular system is a model synapse for the study of synaptic physiology and neurotransmission, and is a valuable research tool that has defined genetics and is accessible to experimental manipulation. Larvae can be dissected to expose the body wall musculature, central nervous system, and peripheral nerves. The muscles of Drosophila and their innervation pattern are well characterized and muscles are easy to access for intracellular recording. Individual muscles can be identified by their location and orientation within the 8 abdominal segments, each with 30 muscles arranged in a pattern that is repeated in segments A2 - A7. Dissected drosophila larvae are thin and individual muscles and bundles of motor neuron axons can be visualized by transillumination1. Transgenic constructs can be used to label target cells for visual identification or for manipulating gene products in specific tissues. In larvae, excitatory junction potentials (EJP&#8217;s) are generated in response to vesicular release of glutamate from the motoneurons at the synapse. In dissected larvae, the EJP can be recorded in the muscle with an intracellular electrode. Action potentials can be artificially evoked in motor neurons that have been cut posterior to the ventral ganglion, drawn into a glass pipette by gentle suction and stimulated with an electrode. These motor neurons have distinct firing thresholds when stimulated, and when they fire simultaneously, they generate a response in the muscle. Signals transmitted across the NMJ synapse can be recorded in the muscles that the motor neurons innervate. The EJP&#8217;s and minature excitatory junction potentials (mEJP&#8217;s) are seen as changes in membrane potential. Electrophysiological responses are recorded at room temperature in modified minimal hemolymph-like solution2 (HL3) that contains 5 mM Mg2+ and 1.5 mM Ca2+. Changes in the amplitude of evoked EJP&#8217;s can indicate differences in synaptic function and structure. Digitized recordings are analyzed for EJP amplitude, mEJP frequency and amplitude, and quantal content.

Here we describe electrophysiological methods for measuring synaptic transmission at the neuromuscular junction of Drosophila larva. Evoked release is initiated artificially by stimulating the motor neuron axons, and transmission through the NMJ can be measured by the postsynaptic response evoked in the muscle.

Drosophila의 애벌레의 NMJ에서 시냅틱 후보를 녹음 Electrophysiological 방법

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. ...

Electrophysiological Recording in the Drosophila Embryo

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite isolated from each other, with largely independent histories and scientific communities. However, the interface between these usually disparate fields is the developmental programs underlying acquisition of functional electrical signaling properties and differentiation of functional chemical synapses during the final phases of neural circuit formation. This interface is a critically important area for investigation. In Drosophila, these phases of functional development occur during a period of &lt;8 hours (at 25&deg;C) during the last third of embryogenesis. This late developmental period was long considered intractable to investigation owing to the deposition of a tough, impermeable epidermal cuticle. A breakthrough advance was the application of water-polymerizing surgical glue that can be locally applied to the cuticle to enable controlled dissection of late-stage embryos. With a dorsal longitudinal incision, the embryo can be laid flat, exposing the ventral nerve cord and body wall musculature to experimental investigation. Whole-cell patch-clamp techniques can then be employed to record from individually-identifiable neurons and somatic muscles. These recording configurations have been used to track the appearance and maturation of ionic currents and action potential propagation in both neurons and muscles. Genetic mutants affecting these electrical properties have been characterized to reveal the molecular composition of ion channels and associated signaling complexes, and to begin exploration of the molecular mechanisms of functional differentiation. A particular focus has been the assembly of synaptic connections, both in the central nervous system and periphery. The glutamatergic neuromuscular junction (NMJ) is most accessible to a combination of optical imaging and electrophysiological recording. A glass suction electrode is used to stimulate the peripheral nerve, with excitatory junction current (EJC) recordings made in the voltage-clamped muscle. This recording configuration has been used to chart the functional differentiation of the synapse, and track the appearance and maturation of presynaptic glutamate release properties. In addition, postsynaptic properties can be assayed independently via iontophoretic or pressure application of glutamate directly to the muscle surface, to measure the appearance and maturation of the glutamate receptor fields. Thus, both pre- and postsynaptic elements can be monitored separately or in combination during embryonic synaptogenesis. This system has been heavily used to isolate and characterize genetic mutants that impair embryonic synapse formation, and thus reveal the molecular mechanisms governing the specification and differentiation of synapse connections and functional synaptic signaling properties.

Electrophysiological Recording in the Drosophila Embryo

Electrophysiological recordings from Drosophila embryos allow analyses of developing muscle and neuron electrical properties, as well as characterization of functional synaptogenesis at the glutamatergic neuromuscular junction and central cholinergic and GABAergic synapses.

에 Electrophysiological 녹음 Drosophila 엠브료

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

의 전자 현미경을위한 배아의 준비 Drosophila 태아의 심장 관

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

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.

Drosophila의 배아들 중에서 - 생체내의 원심 분리

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

Upright Imaging of Drosophila Embryos

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current techniques used to view such processes are somewhat limited. The following protocol describes a new technique for mounting fixed and labeled Drosophila embryos for coronal viewing with confocal imaging. This method consists of embedding embryos between two layers of glycerin jelly mounting media, and imaging jelly strips positioned upright. The first step for sandwiching the embryos is to make a thin bedding of glycerin jelly on a slide. Next, embryos are carefully aligned on this surface and covered with a second layer of jelly. After the second layer is solidified, strips of jelly are cut and flipped upright for imaging. Alternatives are described for visualizing the embryos depending upon the type of microscope stand to be used. Since all cells along the dorsal-ventral axis are imaged within a single confocal Z-plane, our method allows precise measurement and comparison of fluorescent signals without photobleaching or light scattering common to 3D reconstructions of longitudinally mounted embryos.

Upright Imaging of Drosophila Embryos

Here we present a mounting protocol for stained Drosophila embryos in an upright position that allows imaging of cross-sections using Confocal microscopy.

의 입식 이미징 Drosophila 태아

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current ...

Optimized Transfection Strategy for Expression and Electrophysiological Recording of Recombinant Voltage-Gated Ion Channels in HEK-293T Cells

The in vitro expression and electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T) is a ubiquitous research strategy. HEK-293T cells must be plated onto glass coverslips at low enough density so that they are not in contact with each other in order to allow for electrophysiological recording without confounding effects due to contact with adjacent cells. Transfected channels must also express with high efficiency at the plasma membrane for whole-cell patch clamp recording of detectable currents above noise levels. Heterologous ion channels often require long incubation periods at 28&deg;C after transfection in order to achieve adequate membrane expression, but there are increasing losses of cell-coverslip adhesion and membrane stability at this temperature. To circumvent this problem, we developed an optimized strategy to transfect and plate HEK-293T cells. This method requires that cells be transfected at a relatively high confluency, and incubated at 28&deg;C for varying incubation periods post-transfection to allow for adequate ion channel protein expression. Transfected cells are then plated onto glass coverslips and incubated at 37&deg;C for several hours, which allows for rigid cell attachment to the coverslips and membrane restabilization. Cells can be recorded shortly after plating, or can be transferred to 28&deg;C for further incubation. We find that the initial incubation at 28&deg;C, after transfection but before plating, is key for the efficient expression of heterologous ion channels that normally do not express well at the plasma membrane. Positively transfected, cultured cells are identified by co-expressed eGFP or eGFP expressed from a bicistronic vector (e.g. pIRES2-EGFP) containing the recombinant ion channel cDNA just upstream of an internal ribosome entry site and an eGFP coding sequence. Whole-cell patch clamp recording requires specialized equipment, plus the crafting of polished recording electrodes and L-shaped ground electrodes from borosilicate glass. Drug delivery to study the pharmacology of ion channels can be achieved by directly micropipetting drugs into the recording dish, or by using microperfusion or gravity flow systems that produce uninterrupted streams of drug solution over recorded cells.

Reliable method for highly efficient in vitro expression and subsequent electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T).

표현과 HEK - 293T 세포의 재조합 전압 - 게이 티드 이온 채널 Electrophysiological 녹음에 최적화된 Transfection 전략

The in vitro expression and electrophysiological recording of recombinant voltage-gated ion channels in cultured human embryonic kidney cells (HEK-293T) ...

Isolation and Purification of Kinesin from Drosophila Embryos

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a variety of neurodegenerative diseases, understanding microtubule based motor transport and its regulation will likely ultimately lead to improved therapeutic approaches. Kinesin-1 is a eukaryotic motor protein which moves in an anterograde (plus-end) direction along microtubules (MTs), powered by ATP hydrolysis. Here we report a detailed purification protocol to isolate active full length kinesin from Drosophila embryos, thus allowing the combination of Drosophila genetics with single-molecule biophysical studies. Starting with approximately 50 laying cups, with approximately 1000 females per cup, we carried out overnight collections. This provided approximately 10 ml of packed embryos. The embryos were bleach dechorionated (yielding approximately 9 grams of embryos), and then homogenized. After disruption, the homogenate was clarified using a low speed spin followed by a high speed centrifugation. The clarified supernatant was treated with GTP and taxol to polymerize MTs. Kinesin was immobilized on polymerized MTs by adding the ATP analog, 5'-adenylyl imidodiphosphate at room temperature. After kinesin binding, microtubules were sedimented via high speed centrifugation through a sucrose cushion. The microtubule pellet was then re-suspended, and this process was repeated. Finally, ATP was added to release the kinesin from the MTs. High speed centrifugation then spun down the MTs, leaving the kinesin in the supernatant. This kinesin was subjected to a centrifugal filtration using a 100 KD cut off filter for further purification, aliquoted, snap frozen in liquid nitrogen, and stored at -80 &deg;C. SDS gel electrophoresis and western blotting was performed using the purified sample. The motor activity of purified samples before and after the final centrifugal filtration step was evaluated using an in vitro single molecule microtubule assay. The kinesin fractions before and after the centrifugal filtration showed processivity as previously reported in literature. Further experiments are underway to evaluate the interaction between kinesin and other transport related proteins.

Isolation and Purification of Kinesin from Drosophila Embryos

This is a protocol to isolate active full length Kinesin from Drosophila embryos for single-molecule biophysical studies. We show how to collect embryos, make the embryo lysate, and then polymerize microtubules (MTs). Kinesin is purified by immobilizing it on the MTs, spinning down the Kinesin-MT complexes, and then releasing the kinesin from the MTs via ATP addition.

Kinesin의 분리 및 정제에서 Drosophila 태아

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a ...

Preparing Individual Drosophila Egg Chambers for Live Imaging

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, cell differentiation and organogenesis 1. Correct preparation of the experimental samples is a crucial, often neglected, step. The goal of preparation is to ensure physiological relevance and to establish optimal imaging conditions. To maintain tissue viability, it is critical to avoid dehydration, hypoxia, overheating or medium deterioration 2. 
 The Drosophila egg chamber is a well established system for examining questions relating, but not limited, to body patterning, mRNA localization and cytoskeletal organization 3,4. For early- and mid-stage egg chambers, mounting in halocarbon oil is good for survival in that it allows free diffusion of oxygen, prevents dehydration and hypoxia and has superb optical properties for microscopy. Imaging of fluorescent proteins is possible through the introduction of transgenes into the egg chamber or physical injection of labeled RNA, protein or antibodies 5-7. For example, addition of MS2 constructs to the genome of animals enables real time observation of mRNAs in the oocyte 8. These constructs allow for in vivo labeling of mRNA through utilization of the MS2 bacteriophage RNA stem loop interaction with its coat protein 9. 
 Here, we present a protocol for the extraction of ovaries as well as isolating individual ovarioles and egg chambers from the female Drosophila. For a detailed description of Drosophila oogenesis see Allan C. Spradling (1993, reprinted 2009) 10.

Preparing Individual Drosophila Egg Chambers for Live Imaging

The Drosophila egg chamber is an excellent model for studying the mechanisms of mRNA localization. In order to capture the dynamic events that underpin the processes of localization, rapid high resolution imaging of live tissue is required. Here, we present a protocol for dissection and imaging of live samples with minimal disruption.

개별 준비 Drosophila</em라이브 이미징을위한> 계란 회의소

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, ...

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

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

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

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

수확 및 준비 Drosophila 태아