蚊から中腸および唾液腺組織の解離は、1Xリン酸塩の事前準備が固定化° C、まで、4の温度に供することにより生理食塩水（1 × PBS）ソリューションと蚊の麻酔をバッファする必要があります。蚊は、氷の上に冷たい保たれるペトリ皿の中に配置することにより麻酔のまま。必要な他の材料は、次のとおり10倍目標、ピペッター、先の細いピンセット、スライドガラス、針先端のプローブを装着した光学顕微鏡を。 中腸の解剖<ol><li>光学顕微鏡下にマウントされたスライドグラス上に1 × PBSのドロップを置きます。 </li><li>針先端のプローブを用いて蚊胸部を刺すことで、準備したスライド上に蚊を転送する。 </li><li>プローブで蚊を押しながら、最後に腹部の体節から2番目を持ち、ゆっくりとワンタッチで蚊腹部をやってのけるために鉗子を使用する。中腸は固定化胸部に接続されたままにしてください。 </li><li>腹部を捨てる。胸部から腸を切り離すために鉗子を使用してください。 </li></ol>唾液腺解剖<ol><li>光学顕微鏡下にマウントされたスライドグラス上に1 × PBSのドロップを置きます。 </li><li>針先端のプローブを用いて胸部をstabblingで蚊を拾う。 </li><li>あなたの指を使って蚊の足をやってのける。 </li><li>スライドの上に蚊を転送する。 </li><li>鉗子を使用して蚊の頭を削除します。 </li><li>プローブと蚊胸部を押しながら、静かに胸部を下にプッシュする別のプローブを使用。 </li><li>唾液腺は、胸部の前部に位置しており、針先端のプローブを使用し、胸部へのグランドを接続するアタッチメントを切断することによって単離することができる。 2つの横方向の葉と中葉：無傷の唾液腺は3つの葉で構成されています。 </li></ol>

The authors have nothing to disclose.

dissection of midgut and salivary glands from ae. aegypti mosquitoes

蚊の中腸と唾液腺では、マラリア原虫やデング熱ウイルスなどの病原体のための主要な入り口と出口のポイントです。このビデオプロトコルは、ネッタイシマカ蚊から中腸と唾液腺の除去のための解剖のテクニックを示します。

Watch this Scientific Journal Video about Dissection of Midgut and Salivary Glands from Ae. aegypti Mosquitoes at JoVE.com

Dissection of Midgut and Salivary Glands from Ae. aegypti Mosquitoes

蚊の中腸と唾液腺には、熱帯熱マラリア原虫やデング熱ウイルスのようなベクトルの病原体のためのキーエントリーと出口ポイントです。このビデオでは、ネッタイシマカ蚊から中腸と唾液腺を除去するために解剖のテクニックを示します。

AEから中腸と唾液腺の解剖。ネッタイシマカ蚊

The mosquito midgut and salivary glands are key entry and exit points for pathogens such as Plasmodium parasites and Dengue viruses. This video protocol ...

dissection-of-midgut-and-salivary-glands-from-ae-aegypti-mosquitoes

Research

JoVE Journal

Biology

34.7K ビュー.  University of California, Irvine (UCI). 蚊の中腸と唾液腺には、熱帯熱マラリア原虫やデング熱ウイルスのようなベクトルの病原体のためのキーエントリーと出口ポイントです。このビデオでは、ネッタイシマカ蚊から中腸と唾液腺を除去するために解剖のテクニックを示します。

ビデオ: Dissection of Midgut and Salivary Glands from Ae. aegypti Mosquitoes

Microinjection of A. aegypti Embryos to Obtain Transgenic Mosquitoes

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue fever.  The techniques for correctly preparing microinjection needles, dessicating embryos, and performing microinjection are demonstrated.

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue fever. The techniques for correctly preparing microinjection needles, dessicating embryos, and performing microinjection are demonstrated.

トランスジェニック蚊を取得するA.ネッタイシマカの胚のマイクロインジェクション

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue ...

生物学

Protocol for Dengue Infections in Mosquitoes (A. aegypti) and Infection Phenotype Determination

The purpose of this procedure is to infect the Aedes mosquito with dengue virus in a laboratory condition and examine the infection level and dynamic of the virus in the mosquito tissues. This protocol is routinely used for studying mosquito-virus interactions, especially for identification of novel host factors that are able to determine vector competence. The entire experiment must be conducted in a BSL2 laboratory. Similar to Plasmodium falciparum infections, proper attire including gloves and lab coat must be worn at all times. After the experiment, all the materials that came in contact with the virus need to be treated with 75% ethanol and bleached before proceeding with normal washing. All other materials need to be autoclaved before discarding them.

Protocol for Dengue Infections in Mosquitoes A. aegypti and Infection Phenotype Determination

Once a gene is identified as potentially refractory for the dengue virus, it must be evaluated for it's role in preventing viral infections within the mosquito. This protocol illustrates how the extent of dengue infections of mosquitoes can be assayed. The techniques for growing up the virus in culture, membrane feeding mosquitoes human blood, and assaying viral titers in the mosquito midgut are demonstrated.

デング熱蚊の感染症（A.ネッタイシマカ）と感染症の表現型決定のためのプロトコル

The purpose of this procedure is to infect the Aedes mosquito with dengue virus in a laboratory condition and examine the infection level and dynamic of ...

Building a Better Mosquito: Identifying the Genes Enabling Malaria and Dengue Fever Resistance in A. gambiae and A. aegypti Mosquitoes

In this interview, George Dimopoulos focuses on the physiological mechanisms used by mosquitoes to combat Plasmodium falciparum and dengue virus infections.   Explanation is given for how key refractory genes, those genes conferring resistance to vector pathogens, are identified in the mosquito and how this knowledge can be used to generate transgenic mosquitoes that are unable to carry the malaria parasite or dengue virus.

In this interview, George Dimopoulos focuses on the physiological mechanisms used by mosquitoes to combat Plasmodium falciparum and dengue virus infections. Explanation is given for how key refractory genes, those genes conferring resistance to vector pathogens, are identified in the mosquito and how this knowledge can be used to generate transgenic mosquitoes that are unable to carry the malaria parasite or dengue virus.

より良い蚊の構築：A. gambiaeとA.ネッタイシマカ蚊のマラリアとデング熱の抵抗を使用可能にする遺伝子を特定

In this interview, George Dimopoulos focuses on the physiological mechanisms used by mosquitoes to combat Plasmodium falciparum and dengue virus ...

Dissection of Organs from the Adult Zebrafish

Over the last 20 years, the zebrafish has become a powerful model organism for understanding vertebrate development and disease.  Although experimental analysis of the embryo and larva is extensive and the morphology has been well documented, descriptions of adult zebrafish anatomy and studies of development of the adult structures and organs, together with techniques for working with adults are lacking. The organs of the larva undergo significant changes in their overall structure, morphology, and anatomical location during the larval to adult transition.  Externally, the transparent larva develops its characteristic adult striped pigment pattern and paired pelvic fins, while internally, the organs undergo massive growth and remodeling.  In addition, the bipotential gonad primordium develops into either testis or ovary.  This protocol identifies many of the organs of the adult and demonstrates methods for dissection of the brain, gonads, gastrointestinal system, heart, and kidney of the adult zebrafish. The dissected organs can be used for in situ hybridization, immunohistochemistry, histology, RNA extraction, protein analysis, and other molecular techniques.  This protocol will assist in the broadening of studies in the zebrafish to include the remodeling of larval organs, the morphogenesis of organs specific to the adult and other investigations of the adult organ systems.

This protocol describes a procedure for identifying and dissecting organs from the adult zebrafish.

アダルトゼブラフィッシュからの臓器の解剖

Over the last 20 years, the zebrafish has become a powerful model organism for understanding vertebrate development and disease.  Although experimental ...

RNAi-mediated Gene Knockdown and In Vivo Diuresis Assay in Adult Female Aedes aegypti Mosquitoes

This video protocol demonstrates an effective technique to knockdown a particular gene in an insect and conduct a novel bioassay to measure excretion rate. This method can be used to obtain a better understanding of the process of diuresis in insects and is especially useful in the study of diuresis in blood-feeding arthropods that are able to take up huge amounts of liquid in a single blood meal. 
This RNAi-mediated gene knockdown combined with an in vivo diuresis assay was developed by the Hansen lab to study the effects of RNAi-mediated knockdown of aquaporin genes on Aedes aegypti mosquito diuresis1. 
The protocol is setup in two parts: the first demonstration illustrates how to construct a simple mosquito injection device and how to prepare and inject dsRNA into the thorax of mosquitoes for RNAi-mediated gene knockdown. The second demonstration illustrates how to determine excretion rates in mosquitoes using an in vivo bioassay.

RNAi-mediated Gene Knockdown and In Vivo Diuresis Assay in Adult Female Aedes aegypti Mosquitoes

In this protocol we combine RNAi-mediated gene silencing with an in vivo diuresis assay to study the effects knockdown of genes of interest has on mosquito fluid excretion.

RNAiによる遺伝子ノックダウンと<emインビボで&gt;</em大人の女性の&gt;利尿アッセイ<em&gt;ネッタイシマカ</em&gt;蚊

This video protocol demonstrates an effective technique to knockdown a particular gene in an insect and conduct a novel bioassay to measure excretion ...

Maintaining Aedes aegypti Mosquitoes Infected with Wolbachia

Aedes aegypti mosquitoes experimentally infected with Wolbachia are being utilized in programs to control the spread of arboviruses such as dengue, chikungunya and Zika. Wolbachia-infected mosquitoes can be released into the field to either reduce population sizes through incompatible matings or to transform populations with mosquitoes that are refractory to virus transmission. For these strategies to succeed, the mosquitoes released into the field from the laboratory must be competitive with native mosquitoes. However, maintaining mosquitoes in the laboratory can result in inbreeding, genetic drift and laboratory adaptation which can reduce their fitness in the field and may confound the results of experiments. To test the suitability of different Wolbachia infections for deployment in the field, it is necessary to maintain mosquitoes in a controlled laboratory environment across multiple generations. We describe a simple protocol for maintaining Ae. aegypti mosquitoes in the laboratory, which is suitable for both Wolbachia-infected and wild-type mosquitoes. The methods minimize laboratory adaptation and implement outcrossing to increase the relevance of experiments to field mosquitoes. Additionally, colonies are maintained under optimal conditions to maximize their fitness for open field releases.

Maintaining Aedes aegypti Mosquitoes Infected with Wolbachia

Aedes aegypti mosquitoes infected with Wolbachia are being released into natural populations to suppress the transmission of arboviruses. We describe methods to rear Ae. aegypti with Wolbachia infections in the laboratory for experiments and field release, taking precautions to minimize laboratory adaptation and selection.

ネッタイシマカ蚊ボルバキアに感染維持

Aedes aegypti mosquitoes experimentally infected with Wolbachia are being utilized in programs to control the spread of arboviruses such as dengue, ...

Isolation of Myoepithelial Cells from Adult Murine Lacrimal and Submandibular Glands

The lacrimal gland (LG) is an exocrine tubuloacinar gland that secretes an aqueous layer of tear film. The LG epithelial tree is comprised of acinar, ductal epithelial, and myoepithelial cells (MECs). MECs express alpha smooth muscle actin (&#945;SMA) and have a contractile function. They are found in multiple glandular organs and are of ectodermal origin. In addition, the LG contains SMA+ vascular smooth muscle cells of endodermal origin called pericytes: contractile cells that envelop the surface of vascular tubes. A new protocol allows us to isolate both MECs and pericytes from adult murine LGs and submandibular glands (SMGs). The protocol is based on the genetic labeling of MECs and pericytes using the SMACreErt2/+:Rosa26-TdTomatofl/fl mouse strain, followed by preparation of the LG single-cell suspension for fluorescence activated cell sorting (FACS). The protocol allows for the separation of these two cell populations of different origins based on the expression of the epithelial cell adhesion molecule (EpCAM) by MECs, whereas pericytes do not express EpCAM. Isolated cells could be used for cell cultivation or gene expression analysis.

The lacrimal gland (LG) has two cell types expressing &#945;-smooth muscle actin (&#945;SMA): myoepithelial cells (MECs) and pericytes. MECs are of ectodermal origin, found in many glandular tissues, while pericytes are vascular smooth muscle cells of endodermal origin. This protocol isolates MECs and pericytes from murine LGs.

成人ネズミ涙腺および顎下腺からの筋上皮細胞の単離

The lacrimal gland (LG) is an exocrine tubuloacinar gland that secretes an aqueous layer of tear film. The LG epithelial tree is comprised of acinar, ...

Preparing Irradiated and Marked Male Aedes aegypti Mosquitoes for Release in an Operational Sterile Insect Technique Program

The control of such human diseases as dengue, Zika, and chikungunya relies on the control of their vector, the Aedes aegypti mosquito, because there is no prevention. Control of mosquito vectors can rely on chemicals applied to the immature and adult stages, which can contribute to the mortality of non-targets and more importantly, lead to insecticide resistance in the vector. The sterile insect technique (SIT) is a method of controlling populations of pests through the release of sterilized adult males that mate with wild females to produce non-viable offspring. This paper describes the process of producing sterile males for use in an operational SIT program for the control of Aedes aegypti mosquitoes. Outlined here are the steps used in the program including rearing and maintaining a colony, separating male and female pupae, irradiating and marking adult males, and shipping Aedes aegypti males to the release site. Also discussed are procedural caveats, program limitations, and future objectives.

Preparing Irradiated and Marked Male Aedes aegypti Mosquitoes for Release in an Operational Sterile Insect Technique Program

The sterile insect technique (SIT) is used to control specific, medically important mosquito populations that may be resistant to chemical controls. Here, we describe a method of mass rearing and preparation of sterile male mosquitoes for release in an operational SIT program targeting the Aedes aegypti mosquito.

照射され、マークされたオスの ネッタイシマカ を、操作可能な滅菌昆虫技術プログラムで放出するための準備

The control of such human diseases as dengue, Zika, and chikungunya relies on the control of their vector, the Aedes aegypti mosquito, because there is no ...

Immunofluorescent Staining for Visualization of Heterochromatin Associated Proteins in Drosophila Salivary Glands

Visualization of heterochromatin aggregates by immunostaining can be challenging. Many mammalian components of chromatin are conserved in Drosophila melanogaster. Therefore, it is an excellent model to study heterochromatin formation and maintenance. Polytenized cells, such as the ones found in salivary glands of third instar D. melanogaster larvae, provide an excellent tool to observe the chromatin amplified nearly a thousand times and have allowed researchers to study changes in the distribution of heterochromatin in the nucleus. Although the observation of heterochromatin components can be carried out directly in polytene chromosome preparations, the localization of some proteins can be altered by the severity of the treatment. Therefore, the direct visualization of heterochromatin in cells complements this type of study. In this protocol, we describe the immunostaining techniques used for this tissue, the use of secondary fluorescent antibodies, and confocal microscopy to observe these heterochromatin aggregates with greater precision and detail.

Immunofluorescent Staining for Visualization of Heterochromatin Associated Proteins in Drosophila Salivary Glands

This protocol aims to visualize heterochromatin aggregates in Drosophila polytene cells.

ショ ウジョウバエ 唾液腺におけるヘテロクロマチン関連タンパク質の可視化のための免疫蛍光染色

Visualization of heterochromatin aggregates by immunostaining can be challenging. Many mammalian components of chromatin are conserved in Drosophila ...

Using the Fluorescent Dye, Rhodamine B, to Study Mating Competitiveness in Male Aedes aegypti Mosquitoes

The success of sterile or incompatible insect technique-based population suppression programs depends on the ability of released males to compete for wild-type females and induce sterility in the target population. Hence, laboratory assessment of male mating competitiveness is essential for evaluating the release strain&#8217;s fitness before field release. Conventionally, such an assay is performed by determining the proportion of viable eggs produced by the females after being simultaneously exposed to two sets of males (wild-type and release strains) for copulation. However, this process is time-consuming and laborious due to the need to first blood-feed the females for egg production and then hatch and enumerate the hatched eggs to determine egg viability.&#160;
Moreover, this method cannot discern the degree of competitiveness between two sterile or Wolbachia-infected mosquito lines as wild-type female mosquitoes will only produce non-viable eggs upon mating with both. To circumvent these limitations, this paper describes a more direct method of measuring male mosquito mating competitiveness in laboratory settings using the fluorescent dye, rhodamine B (RhB), which can be used to mark males by feeding them in sucrose solution containing RhB. After the mating assay, the presence of fluorescing sperms in the spermathecae of a female can be used to determine her mating partner. This method is cost-effective, reduces the experimental time by 90% and allows comparison of mating fitness between two sterile or Wolbachia-infected lines.&#160;

Using the Fluorescent Dye, Rhodamine B, to Study Mating Competitiveness in Male Aedes aegypti Mosquitoes

Here, we present a protocol for studying mating competitiveness of male Aedes aegypti using fluorescent dye as a marker. Female mosquitoes are exposed to both marked and unmarked males for copulation. Post mating, their spermathecae are examined under a fluorescence microscope to determine their mating partner.

蛍光色素、ローダミンBを用いて、オス のアエデスアエジプティ 蚊の交配競争力を研究する

The success of sterile or incompatible insect technique-based population suppression programs depends on the ability of released males to compete for ...