ベテラン総務省からメリットグラントと国立衛生研究所のNHLBIからT32-HL098062の助成金は、この作品を支持した。我々は感謝ランドルフH·ヘイスティング、医学博士の助言を認めるしたいおよびVAサンディエゴのヘルスケア·システムの獣医療部隊のアドバイスとサポート。

1。動物の準備<ol><li> （小さいマウスが専門家によって挿管することができます）8週間以上経過しており、以上の20グラムのマウスを入手します。 </li><li>麻酔<ol><li>腹腔内麻酔前のようにケタミンおよびキシラジンの20 mg / kgを、それぞれ、をマウスに注射する。 （この用量は完全にマウスを麻酔するには不十分であるが、人工呼吸器に、挿管した後、安全な転送を容易にする。）が、投与量の調整は、制度的に獣医師に相談して麻酔応答に応じて必要な場合があります。） </li><li> 90〜120秒の誘導チャンバ内の3.5％イソフルラン/酸素を用いたマウスに麻酔を誘導する。 </li><li>慎重に呼吸数を観察します。それは、以下45以上BPMに徐々に遅くする必要があります。 </li></ol></li></ol> 2。挿管の準備<ol><li>誘導室からマウスを取り外し、しっかりと首筋。 </li><li> GENTするクッション細かい鉗子を使用してくださいLY口から舌を伸ばす。 </li><li> 2本の指の間に舌を保持し、穏やかな力を加えることにより、舌の拡張を維持する。 </li><li>上下動と耳鏡の鏡の上でマウスを上に引き上げます。 </li><li>しっかりscruffingながらオトスコープの接眼レンズをのぞきながら舌と首筋により穏やかにマウスを引き上げます。 </li><li>声帯を十分に確認してください。彼らは、この時点で簡単に表示されるはずです。喉頭のaditus（喉頭開口部の構造）は、比較的白になります。呼吸のたびにコードの動きがあるはずです。コー​​ドが動いていないか、不十分で可視化されている場合は、わずかに動物を回転させ、ゆっくりと首を過伸展。 </li><li>利き手で、鉛筆のように、カテーテルの先端を通って延びるPE10チューブの長さ1cmで、カテーテル内に20のG、1を保持し、鏡の横に挿入します。 PEチューブはスタイやブジーとして機能します。 </li><li> PE10 tubiに指示NG（スタイレット）声帯を通って、ハブが下顎切歯のレベルになるまでのPEチューブを介して20gのカテーテル（気管内チューブ）を進める。すぐにこのスタイレットを取り外します。 </li><li>そっと鏡の外動物を取り、実験を通して全身麻酔を維持するために継続的な2％のイソフルランで人工呼吸器に動物を配置することにより、チューブの位置を確認します。挿管を確認するため、PEEPトラップを通過する呼気（気泡）可視化する。食道挿管が数泡になるかもしれないが、それは同様に顕著または気管内挿管と同様に一貫性がありません。また、負のたわみ（ 図1）のトレース気道内圧を観察します。これらは、チューブの適切な配置を確認する。あるいは、IVチューブに少量の水を置き、ETチューブに接続します。呼吸2とマウスとの間での動きの確認。食道挿管、Pの場合案内圧力スイッチのトレースが有意に高い圧力と負の偏向を明らかにします。呼気終末CO 2が参考になるが、技術的にはこれは、小さな換気量（〜200μL）指定された非常に困難であろうし、これを行うための機器は非常に高価である。説明した単純な技術は完全に十分であり、はるかに安価である。イソフルラン麻酔が使用できない場合、すべての手順は、全身麻酔のためのキシラジン10-20 mg / kgのケタミンと併用して80〜120 mg / kgのを用いて行うことができる。 </li></ol> 3。代替技術：直接喉頭鏡検査オトスコープの技術では、主に肺に研究物質の直接点眼用に、他の目的のために容易に使用されている。 <ol><li>誘導チャンバ内で90〜120秒間、3.5％で、この手順のためだけではイソフルランで動物に麻酔を誘導する。 </li><li>動物首筋しっかりと頭蓋底と優しく舌を伸ばす細かいピンセットで。 </li><li>優しく舌を保持し、動物がさらにプルアップすることができなくなるまで鏡（未変更）にまっすぐな動物を引っ張る。 </li><li>声帯だけでは、この技術によって、ほとんどの場合、可視化されますが、動物およびHyper-拡張完全なビューにそれらをもたらすために、首を回転させることができます。 </li><li>声門への点滴を含むゲルローディングピペットを進め、流体を植え付ける。鏡を下咽頭とシールを行っているため、マウスはコードを経由しない任意の残液を吸引します。これは簡単にrhonchorousによって検証された流体が肺の中に完全に分散するまで、呼吸時にマウスが、今になり鳴ります。 </li><li>あるいは、液体の後ろの空気塊と流体の50μLを含む0.5ミリリットル注射器にPE10チューブを接続します。 0.5センチメートルのため声帯を注意深くPEチューブを前進させ、徐々に気道内に内容を放電してください。しかし、下気道への時折のトラウマは、Uを発生するサンプルのより良い配達せずに、このテクニックを歌う。 </li></ol>

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著者は、彼らが競合する経済的利益を持っていないことを宣言します。

本報告書では、非外傷性であり、同じ動物で繰り返し使用することができ、マウスを挿管するための簡単​​で信頼性の高い技術が記載されている。この技術は、適度の和を購入することができ、簡単な実験室又は医療​​機器を用いて達成することができる。もともとヘイスティングス4らによって報告された直接喉頭鏡検査技術もまた、様々な目的のために使用することができる?...

実験用マウスは、生​​物学と病理生物学の主要な哺乳類のモデルとして、事実上すべての種に取って代わっています。実験用マウスは、明確かつ広範囲にヒト疾患のモデルとして価値があることが示されている最小の哺乳類の種であり、人間の生物学と疾患の我々の理解の進歩に非常に貴重であることが判明した。短い妊娠期間と実質的により低いコストは、生物医学研究では一般的なツールとして、ヌルおよびトランスジェニックマウスの開発および研究を可能にした。しかし、平均的な実験用マウス（20〜25 g）を大きさが、結果として、一部の研究者は、より大きな哺乳動物種を研究、生理学的または外科的に基づく研究で彼らの研究を制限しています。これらの研究ではマウスを使用する障害は、深い麻酔下で生理学的測定または広範囲の外科的処置を可能に挿管法で遭遇する困難さである。気管切開1は 、標準的なTEとして用いられてきたこのため、技術と必要な控えめなスキルを実行するのが非常に容易するのではなく、挿管のchnique。しかし、気管切開は、慢性または回復手術の研究を助長されていません。従って、それは、急性実験に制限される。気管切開は、炎症や機密生理的反射神経が重要である、研究の交絡変数を指定できます。 当研究室では、他の研究者によって記載されている技術のほとんどを試してみましたが、さまざまな理由で、それらが不十分発見した。気管切開も外傷性であり、出血や気道の炎症を誘発する。ずっとより問題は、それが都合良く繰り返すことができないことである。設備の適度な投資を必要とする多くの比較的非侵襲的な技術が十分に信頼できるものではない。他の技術は、機器が特定のアプリケーションで動作するかどうかを知らなくても、正当化することは困難で高価な装置を必要とする。従って、我々はもはやtは不要非外傷技術の開発が求められてハン特殊な装置の適度な投資は、迅速かつ確実に達成することができる、慢性モデルで繰り返すことができ、多数の動物で使用することができる。ここでは、このような技術を報告する。

<table border="0" cellpadding="0" cellspacing="0" width="706">
	<colgroup>
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	<tbody>
		<tr height="17">
			<td height="17" style="height:17px;width:411px;">Operating Otoscope Head</td>
			<td style="width:144px;">Welch Allyn</td>
			<td align="right" style="width:83px;">21700</td>
			<td style="width:69px;">&#160;$188.98</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Otoscope Handle</td>
			<td>Welch Allyn</td>
			<td align="right">71000</td>
			<td>&#160;$112.20</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Reuseable Speculum</td>
			<td>Welch Allyn</td>
			<td align="right">22002</td>
			<td>&#160;$ 3.98</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Fine Forceps</td>
			<td>Miltex</td>
			<td>18-779</td>
			<td>&#160;$107.18</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Small clamp stand to hold otoscope</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;"></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Optional Equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Isoflurane Vaporizer (multiple circuit with vacuum waste gas handling)</td>
			<td>Summit Medical</td>
			<td></td>
			<td align="right">$3,000</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Flexivent (Animal Ventilator)</td>
			<td>SCIREQ</td>
			<td></td>
			<td align="right">$35,000</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;"></td>
			<td></td>
			<td></td>
			<td></td>
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			<td height="17" style="height:17px;"></td>
			<td></td>
			<td></td>
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			<td height="17" style="height:17px;">Supplies</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Intravenous catheter (20 ga x 1 inch)</td>
			<td>BD</td>
			<td align="right">381233</td>
			<td>&#160;$&#160; 9.62</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Polyethylene tubing (PE10) (0.011 inch I.D., 0.024 inch O.D.) 100 ft</td>
			<td>Intramedic, Clay-Adams</td>
			<td align="right">427401</td>
			<td>&#160;$115.70</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Ketamine 100 mg/ml (10 ml bottle)</td>
			<td>Butler</td>
			<td align="right">23061</td>
			<td>&#160;$10.00</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Xylazine (100 ml bottle)</td>
			<td>Vedco</td>
			<td align="right">24105</td>
			<td>&#160;$20.00</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">Isoflurane (250 ml bottle)</td>
			<td></td>
			<td></td>
			<td>&#160;$15.00</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;">vecuronium bromide 10 mg/10 ml</td>
			<td>Pfizer</td>
			<td>NDC 0069-0094-01</td>
			<td align="right">$15.00</td>
		</tr>
	</tbody>
</table>

endotracheal intubation in mice via direct laryngoscopy using an otoscope

マウスは、野生型およびトランスジェニックの両方が、現在、生物医学研究において主要な哺乳類のモデルである。挿管や人工​​呼吸は深い麻酔や肺機能の測定値の下に手術を必要とする全体の動物実験のために必要である。気管切開は、機械的な換気を可能にするために、これらのマウスでは気道に挿管するための標準となっている。経口気管内挿管が原因かなりの技術的な難しさや専門性の高い高価な装置の必要性が報告されているが、正常に多くの研究で使用されていなかった。ここでは、2.0ミリメートルの鏡を装備オトスコープを使用し、気管内チューブなどの20のG静脈内カテーテルを使用する直接喉頭鏡検査の手法を報告する。私たちは、挿管し、マウスでは肺機能の正確な評価を行うために広くかつ確実に、この技術を使用している。この技術は、経験豊富な手の中に本質的に動物の損失で、安全であることが証明された。さらに、この手法慢性モデルにおけるマウスの反復研究のために使用することができる。

上記の技術を持つ挿管は、信頼性と高速です。気管内チューブの適切な配置は、最も容易にします（ 図1）をトレースする気道内圧に（通常は、PEEPトラップ）で人工呼吸器回路の、負のたわみの水没呼気リムから期限切れのガスバブリングを観察することによって検証される。気道内圧のトレースにマイナスの偏向は最も信頼性が高い。他の呼気人工呼吸器回路2に接続?...

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Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope

私たちは、2.0ミリメートルの鏡と耳鏡を使用して直接喉頭鏡検査を介したマウスでは、気管内挿管のための、シンプルな信頼性が高く、比較的安価な方法を開発した。この技術は、非外傷性であり、慢性の実験において反復測定のために使用することができる。私たちは、優れた気管切開または以前に非外科的手法を報告したために見つける。

マウスの気管内挿管<em&gt;経由</emオトスコープの使用&gt;直接喉頭鏡検査

Mice, both wildtype and transgenic, are the principal mammalian model in biomedical research currently. Intubation and mechanical ventilation are ...

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Research

JoVE Journal

Medicine

Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope

30.9K ビュー.  VA San Diego Healthcare System. 私たちは、2.0ミリメートルの鏡と耳鏡を使用して直接喉頭鏡検査を介したマウスでは、気管内挿管のための、シンプルな信頼性が高く、比較的安価な方法を開発した。この技術は、非外傷性であり、慢性の実験において反復測定のために使用することができる。私たちは、優れた気管切開または以前に非外科的手法を報告したために見つける。 

ビデオ: Endotracheal Intubation in Mice via Direct Laryngoscopy Using an Otoscope

Using Quantitative Real-time PCR to Determine Donor Cell Engraftment in a Competitive Murine Bone Marrow Transplantation Model

Murine bone marrow transplantation models provide an important tool in measuring hematopoietic stem cell (HSC) functions and determining genes/molecules that regulate HSCs. In these transplant model systems, the function of HSCs is determined by the ability of these cells to engraft and reconstitute lethally irradiated recipient mice. Commonly, the donor cell contribution/engraftment is measured by antibodies to donor- specific cell surface proteins using flow cytometry. However, this method heavily depends on the specificity and the ability of the cell surface marker to differentiate donor-derived cells from recipient-originated cells, which may not be available for all mouse strains. Considering the various backgrounds of genetically modified mouse strains in the market, this cell surface/ flow cytometry-based method has significant limitations especially in mouse strains that lack well-defined surface markers to separate donor cells from congenic recipient cells. Here, we reported a PCR-based technique to determine donor cell engraftment/contribution in transplant recipient mice. We transplanted male donor bone marrow HSCs to lethally irradiated congenic female mice. Peripheral blood samples were collected at different time points post transplantation. Bone marrow samples were obtained at the end of the experiments. Genomic DNA was isolated and the Y chromosome specific gene, Zfy1, was amplified using quantitative Real time PCR. The engraftment of male donor-derived cells in the female recipient mice was calculated against standard curve with known percentage of male vs. female DNAs. Bcl2 was used as a reference gene to normalize the total DNA amount. Our data suggested that this approach reliably determines donor cell engraftment and provides a useful, yet simple method in measuring hematopoietic cell reconstitution in murine bone marrow transplantation models. Our method can be routinely performed in most laboratories because no costly equipment such as flow cytometry is required.

Determining donor cell engraftment presents a challenge in mouse bone marrow transplant models that lack well-defined phenotypical markers. We described a methodology to quantify male donor cell engraftment in female transplant recipient mice. This method can be used in all mouse strains for the study of HSC functions.

競争力のあるマウス骨髄移植モデルにおけるドナー細胞の生着を​​決定するために、定量的リアルタイムPCRを用いて

Murine bone marrow transplantation models provide an important tool in measuring hematopoietic stem cell (HSC) functions and determining genes/molecules ...

医学

A Simple Method of Mouse Lung Intubation

A simple procedure to intubate mice for pulmonary function measurements would have several advantages in longitudinal studies with limited numbers or expensive animal. One of the reasons that this is not done more routinely is that it is relatively difficult, despite there being several published studies that describe ways to achieve it. In this paper we demonstrate a procedure that eliminates one of the major hurdles associated with this intubation, that of visualizing the trachea during the entire time of intubation. The approach uses a 0.5 mm fiberoptic light source that serves as an introducer to direct the intubation cannula into the mouse trachea. We show that it is possible to use this procedure to measure lung mechanics in individual mice over a time course of at least several weeks. The technique can be set up with relatively little expense and expertise, and it can be routinely accomplished with relatively little training. This should make it possible for any laboratory to routinely carry out this intubation, thereby allowing longitudinal studies in individual mice, thereby minimizing the number of mice needed and increasing the statistical power by using each mouse as its own control.

This paper describes a striaghforward and efficient method of intubating mice for pulmonary function measurements or pulmonary instillation, that allows the mice to recover and be studied at later times. The procedure involves an inexpensive fiberoptic light source that directly illuminates the trachea.

マウス肺挿管の簡単な方法

A  simple procedure to intubate mice for pulmonary function measurements would  have several advantages in longitudinal studies with limited numbers or  ...

Heterotopic Mucosal Engrafting Procedure for Direct Drug Delivery to the Brain in Mice

Delivery of therapeutics into the brain is impeded by the presence of the blood-brain barrier (BBB) which restricts the passage of polar and high molecular weight compounds from the bloodstream and into brain tissue. Some direct delivery success in humans has been achieved via implantation of transcranial catheters; however this method is highly invasive and associated with numerous complications. A less invasive alternative would be to dose the brain through a surgically implanted, semipermeable membrane such as the nasal mucosa that is used to repair skull base defects following endoscopic transnasal tumor removal surgery in humans. Drug transfer though this membrane would effectively bypass the BBB and diffuse directly into the brain and cerebrospinal fluid. Inspired by this approach, a surgical approach in mice was developed that uses a donor septal mucosal membrane engrafted over an extracranial surgical BBB defect. This model has been shown to effectively allow the passage of high molecular weight compounds into the brain. Since numerous drug candidates are incapable of crossing the BBB, this model is valuable for performing preclinical testing of novel therapies for neurological and psychiatric diseases.

A mouse model of human endoscopic skull base reconstruction has been developed that creates a semipermeable interface between the brain and nose using nasal mucosal grafts. This method allows researchers to study delivery to the central nervous system of high molecular weight therapeutics which are otherwise excluded by the blood-brain barrier when administered systemically.

マウスの脳に直接薬剤送達のための異粘膜移植手順

Delivery of therapeutics into the brain is impeded by the presence of the blood-brain barrier (BBB) which restricts the passage of polar and high ...

An Improved Method for Rapid Intubation of the Trachea in Mice

Despite some anatomical and physiological differences, mouse models continue to be an essential tool for studying human lung disease. Bleomycin toxicity is a commonly used model to study both acute lung injury and fibrosis, and multiple methods have been developed for administering bleomycin (and other toxic agents) into the lungs. However, many of these approaches, such as transtracheal instillation, have inherent drawbacks, including the need for strong anesthetics and survival surgery. This paper reports a quick, reproducible method of intratracheal intubation that involves mild inhaled anesthesia, visualization of the trachea, and the use of a surrogate spirometer to confirm exposure. As a proof of concept, 8-12 week old C57BL/6 mice were administered either 2.0 U/kg of bleomycin or an equivalent volume of PBS, and both damage and fibrotic endpoints were measured post-exposure. This procedure allows researchers to treat a large cohort of mice in a relatively short period with little expense and minimal post-procedure care.

This article presents a rapid and simple method for administering bleomycin directly into the mouse trachea via intubation. Key advantages of this method are that it is highly reproducible, easy to master, and does not require specialized equipment or lengthy recovery times.

マウスの気管の迅速な挿管のための改善された方法

Despite some anatomical and physiological differences, mouse models continue to be an essential tool for studying human lung disease. Bleomycin toxicity ...

Inducing Acute Lung Injury in Mice by Direct Intratracheal Lipopolysaccharide Instillation

Airway administration of lipopolysaccharide (LPS) is a common way to study pulmonary inflammation and acute lung injury (ALI) in small animal models. Various approaches have been described, such as the inhalation of aerosolized LPS as well as nasal or intratracheal instillation. The presented protocol describes a detailed step-by-step procedure to induce ALI in mice by direct intratracheal LPS instillation and perform FACS analysis of blood samples, bronchoalveolar lavage (BAL) fluid, and lung tissue. After intraperitoneal sedation, the trachea is exposed and LPS is administered via a 22 G venous catheter. A robust and reproducible inflammatory reaction with leukocyte invasion, upregulation of proinflammatory cytokines, and disruption of the alveolo-capillary barrier is induced within hours to days, depending on the LPS dosage used. Collection of blood samples, BAL fluid, and lung harvesting, as well as the processing for FACS analysis, are described in detail in the protocol. Although the use of the sterile LPS is not suitable to study pharmacologic interventions in infectious diseases, the described approach offers minimal invasiveness, simple handling, and good reproducibility to answer mechanistic immunological questions. Furthermore, dose titration as well as the use of alternative LPS preparations or mouse strains allow modulation of the clinical effects, which can exhibit different degrees of ALI severity or early vs. late onset of disease symptoms.

Presented here is a step-by-step procedure to induce acute lung injury in mice by direct intratracheal lipopolysaccharide instillation and to perform FACS analysis of blood samples, bronchoalveolar lavage fluid, and lung tissue. Minimal invasiveness, simple handling, good reproducibility, and titration of disease severity are advantages of this approach.

直接気管内リポ多糖刺激によるマウスの急性肺損傷の誘導

Airway administration of lipopolysaccharide (LPS) is a common way to study pulmonary inflammation and acute lung injury (ALI) in small animal models. ...

Repeated Orotracheal Intubation in Mice

The literature describes several methods for mouse intubation that either require visualization of the glottis through the oral cavity or incision in the ventral neck for direct confirmation of cannula placement in the trachea. The relative difficulty or the tissue trauma induced to the subject by such procedures can be an impediment to an investigator&#8217;s ability to perform longitudinal studies. This article illustrates a technique in which physical manipulation of the mouse following the use of a depilatory to remove hair from the ventral neck permits transcutaneous visualization of the trachea for orotracheal intubation regardless of degree of skin pigmentation. This method is innocuous to the subject and easily achieved with a limited understanding of murine anatomy. This refined approach facilitates repeated intubation, which may be necessary for monitoring progression of disease or instillation of treatments. Using this method may result in a reduction of the number of animals and technical skill required to measure lung function in mouse models of respiratory disease.

The goal of this article is to describe a refined method of intubation of the laboratory mouse. The method is noninvasive and, therefore, ideal for studies that require serial monitoring of respiratory function and/or instillation of treatments into the lung.

マウスにおける反復性の耳管挿管

The literature describes several methods for mouse intubation that either require visualization of the glottis through the oral cavity or incision in the ...

Blind Endotracheal Intubation in Neonatal Rabbits

The newborn rabbit is a useful animal model for various pathologies and procedures. Airway management of the rabbit is complex due to its anatomical characteristics, which is further complicated in the case of the newborn. Of the different methods of advanced airway management, endotracheal intubation is less aggressive than tracheostomy, and is more feasible than supraglottic management given the lack of supraglottic devices of such a small size. As direct glottis visualization is very difficult in animals this size, this blind intubation model is presented as an effective alternative, especially for experiments requiring prolonged anesthesia. Using this method, we performed blind intubations with a 90% success rate.

We describe a technique of endotracheal intubation in newborn rabbits after esophageal catheterization with a gastric tube.

新生児ウサギにおける盲目の気管内挿管

The newborn rabbit is a useful animal model for various pathologies and procedures. Airway management of the rabbit is complex due to its anatomical ...

Direct Drug Delivery to Kidney via the Renal Artery

There is a need for directed injections to enable increased and specific renal exposure for efficient evaluation of drug targets in the renal research field. Accumulation of drugs in certain organs may give rise to adverse and unwanted effects, depending on the nature of injectate. To minimize spillover and/or accumulation in other tissues, the herein described method directs the formulation into the renal artery bloodstream by inserting a catheter in the infra renal aorta, just below where it branches into the renal artery, resulting in the kidney as first reached organ and distributing of formulation throughout the kidney.
This manuscript provides a detailed description of the method, as well as its challenges and difficulties. It guides the experimenter to become skillful with this type of microsurgery that requires accuracy under sterile conditions. Speed is crucial for minimizing the ischemia and practicing the procedure will increase the chance of successful injections without adverse effects. By modulating the time between injection and reperfusion as well as the injected volume, the risk of spillover to other organs is mitigated. 
Note that this technique is suitable for single dosing strategies.

This manuscript describes a method for targeted delivery to a single kidney via a catheter placed in the infrarenal abdominal aorta in the mouse.

腎動脈を介した腎臓への直接薬物送達

There is a need for directed injections to enable increased and specific renal exposure for efficient evaluation of drug targets in the renal research ...

Endotracheal Intubation Using a Flexible Intubation Endoscope as a Standardized Model for Safe Airway Management in Swine

Endotracheal intubation is often a basic requirement for translational research in porcine models for various interventions that require a secured airway or high ventilation pressures. Endotracheal intubation is a challenging skill,&#160;requiring a minimum number of successful endotracheal intubations to achieve a high success rate under optimal conditions, which is often unachievable for non-anaesthesiology researchers. Due to the specific porcine airway anatomy, a difficult airway can usually be assumed. The impossibility of establishing a secure airway can result in injury, adverse events, or death of the laboratory animal. Using a prospective, randomized, controlled evaluation approach, it has been shown that fiberoptic-assisted endotracheal intubation takes longer but has a higher first-pass success rate than conventional intubation without causing clinically relevant drops in oxygen saturation. This model presents a standardized regimen for endoscopically guided endotracheal intubation, providing a secured airway, especially for researchers who are inexperienced in the technique of endotracheal intubation via direct laryngoscopy. This procedure is expected to minimize animal suffering and unnecessary animal losses.

The use of pigs in research has increased in recent years. Nevertheless, pigs are characterized by difficult airway anatomy. By demonstrating how to perform endoscopically guided endotracheal intubation, the present protocol aims to further increase laboratory animals' safety to avoid animal suffering and unnecessary death.

ブタの安全な気道管理のための標準化モデルとしての柔軟な挿管内内視鏡を使用した気管内挿管

Endotracheal intubation is often a basic requirement for translational research in porcine models for various interventions that require a secured airway ...

<meta charset="utf8"></head><body>非生存ラットモデルのための気管切開術に基づく気管内挿管プロトコル

Endotracheal Intubation via Tracheotomy and Subsequent Thoracotomy in Rats for Non-Survival Applications

Endotracheal intubation and subsequent ventilation are often basic requirements for translational research in rat models for various interventions that require controlled or high ventilation pressures or access to the thoracic cavity and organs. Conventional endoorotracheal intubation using the anatomically existing route through the mouth is well suited for survival experiments. However, this procedure poses some challenges, including generally higher levels of the required experience and technical skill, more advanced equipment, and greater time effort with relevant intubation failure rates and complications such as tracheal perforation, temporary systemic hypooxygenation, and relevant aerial leakage.
This manuscript, therefore, presents a detailed step-by-step protocol for endotracheal intubation through tracheotomy in non-survival rat models when guaranteed intubation success, constant oxygenation levels, high ventilation pressures, or open thoracotomy are required.
The protocol emphasizes the importance of meticulous surgical technique to ensure consistent and reliable outcomes, especially for researchers who are inexperienced or lack routine in the technique of endoorotracheal intubation via direct laryngoscopy. This procedure is, therefore, expected to minimize animal suffering and unnecessary animal losses.

<meta charset="utf8"></head><body>非生存アプリケーションのためのラットにおける気管切開術およびその後の開胸術による気管内挿管術

Here we introduce a standardized procedure for endotracheal intubation via tracheotomy followed by thoracotomy in rats, aimed at enhancing the precision and reproducibility of non-survival applications requiring invasive ventilation and exposure of thoracic organs in in vivo rat models.

非生存アプリケーションのためのラットにおける気管切開術およびその後の開胸術による気管内挿管術

Endotracheal intubation and subsequent ventilation are often basic requirements for translational research in rat models for various interventions that ...