Este trabalho foi generosamente apoiado pelos Institutos Nacionais de Saúde / Instituto Nacional de Distúrbios Neurológicos e Derrame (NIH / NINDS) concessão 1R01NS086981 eo Hilton Fundação Conrad N.. Agradecemos ao Dr. Laurent Bentolila e Dr. Matthew Schibler por sua inestimável ajuda com microscopia confocal. Agradecemos a todos os que contribuíram para o fórum CLAREZA (http://forum.claritytechniques.org). Agradecemos especialmente Dr. Karl Deisseroth para a abertura de seu laboratório para ensinar essa técnica fascinante. Os autores são gratos pelo apoio generoso da Organização Mapeamento Cerebral investigação médica, fundação Mapeamento Cerebral Suporte, Pierson-Lovelace Foundation, a Fundação Ahmanson, Capital Group Companies Charitable Foundation, William M. e Linda R. Fund Filantrópica Dietel e Fundo Northstar . Pesquisa relatada nesta publicação também foi parcialmente apoiado pelo Centro Nacional de Pesquisa de Recursos e pelo Escritório do Diretor da Naçãoal Institutes of Health, com os números premiados C06RR012169, C06RR015431 e S10OD011939. O conteúdo é da exclusiva responsabilidade dos autores e não representam necessariamente as opiniões oficiais dos Institutos Nacionais de Saúde.

Todos os experimentos foram realizados em conformidade com o Animal Care Institucional e diretrizes Use Committee (IACUC). THY1-YFP, PLP-EGFP e PV-TdTomato ratos foram usados ​​para estas experiências, mas nenhum camundongos que expressam proteínas fluorescentes pode ser apagada e fotografada com sucesso. 1. Preparação do Tecido Cuidado: paraformaldeído (PFA) e acrilamida são tóxicos e irritantes. Realizar experimentos utilizando estes reagentes em um exaustor e com equipamento adequado de proteção individual (jaleco, luvas e proteção para os olhos). <ol><li> Sacrificar o animal na câmara de eutanásia com ~ 1 ml de isoflurano até que a respiração cessa (~ 2-3 min). </li><li> Usar uma bomba peristáltica ajustado para ~ 5 ml / min para perfundir intracardial o rato com uma solução de perfusão (Tabela 1), a 4 ° C até que o sangue é limpo a partir de tecidos (5 - 10 min). </li><li> Alterar o perfusato a solução fixadora (Tabela 1)a 4 ° C até que o pescoço e a cauda endureceram significativamente (5-10 min). </li><li> Terminar a perfusão e decapitar o animal. Cuidadosamente dissecar o cérebro do crânio removendo primeiro o tecido conjuntivo em torno do crânio e em seguida a remoção de osso a partir da superfície ventral do crânio e o levantamento do cérebro para fora (Figura 1a), tal como descrito em 20. Nota: Dê um cuidado especial para a remoção de osso que circunda os lobos flocculonodular do cerebelo e os bulbos olfatórios. <ol><li> Dissecar a medula espinal a partir da coluna vertebral através da inserção suavemente a ponta de uma fina, tesoura afiada entre a medula espinal e a coluna vertebral e o corte dos segmentos de coluna vertebral individuais embora, tal como descrito em 21. </li></ol></li><li> Hemisect o cérebro, colocando-o numa matriz de plástico de cérebro de ratinho e de corte ao longo da linha média com uma lâmina de matriz. </li><li> Adicionar cada hemisfério e da medula espinhal para separar 50 ml banheira de centrífugaES com ~ 40 ml de solução de hidrogel (Tabela 1). A partir deste ponto tampa para a frente os tubos com folha de alumínio para proteger os fluoróforos durante todas as etapas subseqüentes. </li><li> Incubar o tubo que contém a amostra em hidrogel a 4 ° C com agitação suave (~ 30 rpm) durante 7 dias. </li></ol> 2. Polimerização <ol><li> Descartar ~ 25 ml de hidrogel, preservando a amostra e ~ 25 ml de hidrogel no tubo de centrifugação. </li><li> Desoxigena-se a amostra removendo a tampa e a colocação do tubo de centrífuga frasco em exsicador e ligar o frasco a vácuo. Aplicar vácuo durante pelo menos 10 minutos para permitir que os gases dissolvidos a sair da solução e formar bolhas. Agite suavemente para desalojar as bolhas. </li><li> Substituir o vácuo no frasco exsicador com gás de azoto a 100% durante 2-3 min. Uma vez que a pressão equaliza (uma vez que a parte superior do frasco exsicador pode ser aberto), rapidamente tampa do tubo para evitar a reintrodução de oxigénio. </li><li> Polímeroize o hidrogel através da colocação do tubo com a amostra num banho de água a 37 ° C durante 3 h até que a polimerização esteja completa. Hidrogel polimerizado irá assumir uma consistência (Figura 1b) do tipo gel. Nota: Uma pequena quantidade de hidrogel não polimerizada no topo é aceitável. </li><li> Usar uma espátula para remover a amostra polimerizada a partir do tubo de centrifugação, depois cortar o excesso de hidrogel a partir da amostra. </li><li> Remover o hidrogel restante, colocando a amostra num fiapos e limpar esfregando suavemente e rolando o tecido nele, deixando para trás apenas uma fina camada de hidrogel polimerizado na amostra. </li></ol> 3. Lipid Remoção Nota: tecido limpou-Lipid é frágil, punho com cuidado. Use um filtro de células durante a drenagem do tubo para evitar a perda de amostra. Além disso, a amostra irá inchar durante o processo de limpeza, no entanto, o inchaço pode ser invertida durante o processo de harmonização do índice de refracção. Cuidado: SodiUM sulfato de dodecilo (SDS) e ácido bórico são irritantes. Realizar experimentos utilizando-os em um exaustor e com equipamento adequado de proteção individual (jaleco, luvas e proteção para os olhos). <ol><li> Transferir a amostra polimerizada para um tubo de centrífuga de 50 ml limpos e adicionar 45 ml de tampão de compensação (Tabela 1). Incubar a 45 ° C com agitação suave (~ 30 rpm). Alterar o tampão por dia durante um período inicial de 7 dias por vazamento fora do buffer de compensação usado para resíduos químicos e adicionando ~ 45 ml de tampão de compensação fresco. Mantenha os tubos de centrífuga contendo as amostras cobertas de folha de alumínio para proteger os fluoróforos. </li><li> Após os primeiros 7 dias, incubar a amostra a C e troca de compensação tampão 40 ° a cada 2 - 3 dias. Continue compensação até que todos os lipídios são dissolvidos e tecido atinge a transparência (Figura 1c). Isto vai ser de 4 - 6 semanas para um hemisfério cerebral do rato e 2 - 4 semanas para uma medula espinal. </li><li> Uma vez que o tecido é limpo, transferiramostra para um novo tubo de centrífuga de 50 ml e lava-se 4 vezes em PBST (Tabela 1), a 40 ° C com agitação suave (~ 30 rpm) durante 24 horas para remover o SDS residual. </li></ol> 4. Coloração Molecular Cuidado: 4 &#39;, 6-diamidino-2-phenylindole Dihydrochloride (DAPI) é um irritante e quaisquer experiências utilizando deve ser realizada em um exaustor e com equipamento adequado de proteção individual (jaleco, luvas e proteção para os olhos). <ol><li> Transferir a amostra para um tubo de centrífuga de 15 ml limpos e encher-se com 1 ug / ml de DAPI em PBST 1x, pH 7,5. Incubar à temperatura ambiente (TA) durante 24 horas. Mantenha os tubos de centrífuga contendo as amostras cobertas de folha de alumínio para proteger os fluoróforos. </li><li> Lavar 3 vezes em PBST à temperatura ambiente durante pelo menos 6 h / lavagem. </li></ol> 5. Índice de Refração Matching Cuidado: 2,2? Tiodietanol (TDE) é um irritante e quaisquer experimentos utilizando-should ser realizada em um exaustor e com equipamento adequado de proteção individual (jaleco, luvas e proteção para os olhos). <ol><li> Transferir a amostra para um tubo de centrífuga de 15 ml limpos e encher-se com 30% (v / v) em PBS 1x TDE, pH 7,5. Incubar a 40 ° C com agitação suave (~ 30 rpm) durante 24 h. </li><li> Troca TDE 30% com 63% (v / v) em PBS 1x TDE, pH 7,5. Incubar a 40 ° C com agitação suave durante 24 horas. Nota: A amostra vai encolher volta ao tamanho de aproximadamente originais. </li><li> Sobre uma superfície plana, limpo rolar um pedaço de adesivo reutilizável para um cilindro com diâmetro uniforme apenas ligeiramente maior do que a espessura da amostra. </li><li> Colocar o cilindro de adesivo reutilizável em um círculo aberto (~ 25 mm de diâmetro interno com uma pequena abertura na parte superior) em um 40 mm prato de vidro de fundo. Se correcções devem ser feitas, remover o cilindro a partir do vidro e cortado com uma lâmina de barbear. </li><li> Utilizar uma ponteira P1000 pipeta para criar um selo entre o adesivo reutilizável e vidro porrolando-o em torno do rebordo exterior do cilindro. </li><li> Lugar ~ 100 mL de 63% TDE para o prato de vidro e se espalhou para cobrir a superfície de vidro dentro do círculo de adesivo reutilizável. </li><li> Usando uma espátula, coloque cuidadosamente a amostra no meio do círculo, tomando cuidado para não formar bolhas. </li><li> Pipeta mais 100 ul de 63% TDE diretamente sobre a amostra, em seguida, coloque imediatamente a 40 milímetros circular tampa de vidro sobre o adesivo reutilizável. </li><li> Usando o indicador, médio e anelar de ambas as mãos, pressione cuidadosamente ao redor do perímetro do círculo até que a tampa de vidro fez contato com a amostra. </li><li> Lentamente trazer o fundo da placa de vidro para uma posição vertical de descanso de encontro a uma superfície, em seguida, adicionar 63% TDE com uma pipeta até que a solução atinge a pequena abertura na parte superior. Usar um cotão livre limpe para secar a ponta da pipeta de modo a que a solução não goteje sobre o vidro. Para evitar bolhas na câmara, não esvaziar a pipeta de todo o caminho. </li><li> Paraa pequena abertura, adicionar de elastómero de silicone para delimitar o círculo e criar uma câmara fechada. Espere 5 minutos para secar antes de imagem. </li></ol> 6. Microscopia Cuidado: Os lasers usados ​​em confocal, iluminação plano único e microscopia folha de luz são muito poderoso e pode causar cegueira. Quaisquer experiências utilizando lasers deve ser realizada após o treinamento extenso e com grande cuidado e proteção para os olhos adequada. <ol><li> Coloque a câmara de imagem com a amostra dentro no palco de um microscópio de varredura a laser confocal. </li><li> Abra o software de imagem (ver Tabela de Materiais). Para ligar o laser de argônio de excitação, clique na guia de configuração na parte superior do programa, em seguida, no ícone do laser. Verifique a caixa ao lado de argônio para energizar o laser e mover a escala deslizante para 30%. Observar o ponteiro lentamente aquecer para a posição seleccionada. <ol><li> Retornar à guia adquirir no topo. Na caixa grande &quot;Configurações caminho do feixe &amp;# 34; ir para a carga / salvar caixa de configuração e selecione o menu drop-down para selecionar um canal de comprimento de onda adequado para a imagem YFP (527 nm). </li></ol></li><li> Selecione objectivos adequados para imagiologia por localizar a hiperligação vermelho escrito &quot;objetivo: objeto atual&quot; abaixo da caixa de configurações do canal. Ao clicar na guia abre todos os objetivos disponíveis e uma selecção irá girar automaticamente a torre. <ol><li> Utilização objectivos (por ex., 10X), com um raio grande de trabalho (maior do que a espessura do tecido a ser visualizado, por exemplo: 11 mm) e uma grande abertura numérica (por ex., 0,3) para maximizar no plano resolução de imagem e minimizar espessura de corte óptico. </li></ol></li><li> Na coluna à esquerda da menus drop-down, abra o &quot;XY: resolução&quot; janela para definir matriz de aquisição, chamado de &quot;formato&quot;, a 1.024 x 1.024 ou um valor adequado para o limite de resolução lateral do objectivo. Defina o fator de zoom mais baixo possível (por exemplo., 10,7). </li><li> Na mesma janela, jogo 8 média de linha e um quadro de parâmetros médios por cair para baixo os menus marcados individualmente em conformidade. Use valores grandes para uma alta relação sinal-ruído e valores pequenos para a aquisição rápida. Nota: 8 médias de linha e uma média quadro vai adquirir imagens de um hemisfério do cérebro do rato de alta qualidade em aproximadamente 4 horas. </li><li> Localize a amostra usando os controles do estágio. Uma vez que um campo representativo é visível, definir o ganho e offset. Nota: O ganho e offset variará substancialmente com base no tipo de tecido, fluoróforos, e / ou característica anatômica a ser trabalhada. Para YFP, definir valores de 760-780 para o ganho e de -1,0 para -1,5 para o deslocamento. </li><li> Selecione &quot;varredura telha&quot; e selecione os limites da região de interesse. Definir os limites superior e inferior do z-stack a ser adquirido. Definir a espessura da secção óptica baseado em secção de limite de resolução óptica do objectivo. Nota: Um objetivo dolimite de resolução óptica secção é definida principalmente pela sua abertura numérica. A maioria dos softwares de controle microscópio irá calcular automaticamente um valor apropriado para o objetivo que está sendo usado. Para uma objectiva 10X com uma abertura numérica de 0,3 que seria de aproximadamente 11 uM. </li><li> Iniciar a aquisição. Nota: Isso pode demorar vários horas. </li><li> Coletar imagens de alta resolução usando objectivos maior ampliação de regiões específicas de interesse. </li></ol>

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The authors have nothing to disclose.

clearing passiva de tecidos de hidrogel incorporado (clareza passiva) é um método simples e barato para limpar grandes pedaços de tecido. Esta abordagem não requer equipamento específico e pode ser facilmente realizada num agitador com temperatura controlada. Durante o período de algumas semanas, mesmo grandes, tecidos altamente mielinizados, como um cérebro inteiro ou medula espinhal, vai se tornar transparente e adequado para microscopia. Embora este relatório concentrou-se a limpeza de tecidos do SNC, Clareza...

The ability to image complete neuroanatomical structures is immensely valuable for understanding the brain in health and disease. Traditionally, 3D imaging has required tissue sectioning to provide the axial resolution and to visualize deep anatomical structures. This approach can produce high-resolution data sets, but requires sophisticated image reconstruction techniques and is very labor intensive. As a result, it has been limited to the imaging of small volumes of tissue1-3. Optical sectioning, on the other hand, is well suited for the creation of high-resolution 3D images of fluorescently labeled tissues. Since optical sectioning is inherently three dimensional, it does not require extensive computation to produce a 3D image volume. However, light scattering and tissue opacity limit the depth at which tissues can be optically sectioned. The depth of imaging is limited to about 150 &#181;m in laser scanning confocal microscopy and to less than 800 &#181;m using two-photon excitation microscopy4-8.
In order to overcome these limitations, several optical clearing techniques have been recently developed and then further refined to permit the deep microscopic imaging of intact tissues. The use of benzyl alcohol and benzyl benzoate (BABB) to render fixed tissues transparent was among one of the earliest approaches9. However, this approach was limited by the quenching of fluorescence in the samples and incomplete clearing of highly myelinated structures10. Refinements of this technique, such as 3D Imaging of Solvent Cleared Organs (3DISCO), have led to very rapid and complete tissue clearing, but still suffer from rapid loss of fluorescent signals, especially yellow fluorescent protein (YFP)10. Water-based clearing solutions, such as Sca/e11 and SeeDB12, preserve fluorescent signals, but do not completely clear highly myelinated tissues. Clear, Unobstructed Brain Imaging Cocktails and Computational analysis (CUBIC) is a promising new optical clearing technology that appears to overcome the limitations of previously developed water-based clearing solutions13. In contrast with other optical clearing techniques, Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY)14 embeds the brain in a porous matrix that provides structural integrity to proteins, nucleic acids and small molecules, while leaving lipids unbound. The lipids are then removed electrophoretically, culminating in an optically cleared brain that can be easily visualized and equally easily probed using commonly available techniques.
Electrophoretic tissue clearing (ETC) is used to remove lipids from hydrogel embedded tissues in CLARITY, however, ETC can be difficult to implement consistently and tissues subject to electrophoresis can exhibit tissue distortion, browning, loss of fluorescence and antibody reactivity. Passive lipid clearing avoids these limitations, but requires increased clearing times15-17. Passive clearing also permits the clearing of large numbers of samples in parallel, since it is not limited by the number of ETC apparatuses available. Decreasing the concentration of paraformaldehyde and excluding bis-acrylamide from the hydrogel have led to great increases in the speed of clearing at the expense of greater tissue expansion16. Less expensive alternatives to the refractive index matching solution used in the original CLARITY technique have been developed17-19. 2,2'-thiodiethanol (TDE) is an inexpensive and rapid brain clearing agent that reverses some of the tissue expansion that occurs during lipid removal18. This report incorporates passive clearing of hydrogel embedded tissue and the use of TDE as a refractive index matching solution to the original CLARITY technique to produce a highly reproducible and inexpensive protocol for the optical clearing of the mouse central nervous system.

<table><tbody><tr><td>10x phosphate buffered saline (PBS)</td><td>Fisher</td><td>BP399-1</td><td>buffers</td></tr><tr><td>32% paraformaldehyde (PFA)</td><td>Electron Microscopy Sciences</td><td>15714-5</td><td>perfusion and hydrogel</td></tr><tr><td>2,2&#039;-thiodiethanol (TDE)</td><td>Sigma-Aldrich</td><td>166782-500G</td><td>refractive index matching solution</td></tr><tr><td>40% acrylamide</td><td>Bio-Rad</td><td>161-0140</td><td>hydrogel</td></tr><tr><td>2% bis-acrylamide</td><td>Bio-Rad</td><td>161-0142</td><td>hydrogel</td></tr><tr><td>VA-044 initiator</td><td>Wako Pure Chemical Industries, Ltd.</td><td>VA044</td><td>hydrogel</td></tr><tr><td>Boric acid</td><td>Sigma-Aldrich</td><td>B7901</td><td>clearing buffer</td></tr><tr><td>Sodium dodecyl sulfate (SDS)</td><td>Fisher</td><td>BP166-5</td><td>clearing buffer</td></tr><tr><td>Sodium hydroxide</td><td>Fisher</td><td>55255-1</td><td>buffers</td></tr><tr><td>Sodium azide</td><td>Sigma-Aldrich</td><td>52002-100G</td><td>preservative</td></tr><tr><td>Triton X-100</td><td>Sigma-Aldrich</td><td>X100-500G</td><td>buffers</td></tr><tr><td>Heparin</td><td>Sigma-Aldrich</td><td>H3393-50KU</td><td>perfusion</td></tr><tr><td>Sodium nitrate</td><td>Fisher</td><td>BP360-500</td><td>perfusion</td></tr><tr><td>DAPI</td><td>Molecular Probes</td><td>D1306</td><td>nuclear stain</td></tr><tr><td>99.9% isoflurane</td><td>Phoenix</td><td>57319-559-06</td><td>anesthetic</td></tr><tr><td>Hydrocholoric acid</td><td>Fisher</td><td>A1445-500</td><td>buffers</td></tr><tr><td>Glass bottom dish</td><td>Willco</td><td>HBSB-5040</td><td>Willco dishes</td></tr><tr><td>Reusable adhesive</td><td>Bostick</td><td>371351</td><td>Blu-Tack</td></tr><tr><td>Silicon elastomer</td><td>World Precision Instruments</td><td>KWIK-CAST</td><td>Kwik-Cast</td></tr><tr><td>Lint-free wipe</td><td>Kimberly-Clark</td><td>34120</td><td>KimWipe</td></tr><tr><td>Mouse brain matrix</td><td>Roboz</td><td>SA-2175</td><td>sectioning tissue</td></tr><tr><td>Matrix blades</td><td>Roboz</td><td>RS-9887</td><td>sectioning tissue</td></tr><tr><td>Peristaltic pump</td><td>Cole-Parmer</td><td>77122-24</td><td>pefusion</td></tr><tr><td>Laser scanning confocal microscope</td><td>Leica</td><td>SP5</td><td>microscopy</td></tr><tr><td>Imaging software</td><td>Leica</td><td>LAS AF</td><td>microscopy</td></tr></tbody></table>

optical clearing of the mouse central nervous system using passive clarity

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique non-linear deformations, dramatically hampering one's ability to evaluate cellular morphology, distribution and connectivity in the central nervous system (CNS). However, optical clearing techniques are changing the way tissues are visualized. These approaches permit one to probe deeply into intact organ preparations, providing tremendous insight into the structural organization of tissues in health and disease. Techniques such as Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) achieve this goal by providing a matrix that binds important biomolecules while permitting light-scattering lipids to freely diffuse out. Lipid removal, followed by refractive index matching, renders the tissue transparent and readily imaged in 3 dimensions (3D). Nevertheless, the electrophoretic tissue clearing (ETC) used in the original CLARITY protocol can be challenging to implement successfully and the use of a proprietary refraction index matching solution makes it expensive to use the technique routinely. This report demonstrates the implementation of a simple and inexpensive optical clearing protocol that combines passive CLARITY for improved tissue integrity and 2,2&#8242;-thiodiethanol (TDE), a previously described refractive index matching solution.

Visualizando a organização estrutural do cérebro é fundamental para a compreensão de como morfologia e conectividade afetar a função cerebral na saúde e na doença. técnicas de limpeza ópticas tornam possível a populações de células de imagem em 3D em tecidos intactos, o que nos permite estudar a morfologia e conectividade coesa. Os ratinhos que expressam proteínas fluorescentes impulsionado por promotores espec...

Watch this Scientific Journal Video about Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY at JoVE.com

Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY

Optical clearing techniques are revolutionizing the way tissues are visualized. In this report we describe modifications of the original Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) protocol that yields more consistent and less expensive results.

Clearing óptica do Sistema Nervoso Central Rato Usando a claridade passiva

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique ...

optical-clearing-mouse-central-nervous-system-using-passive

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

13.4K visualizações.  University of California, Los Angeles. Optical clearing techniques are revolutionizing the way tissues are visualized. In this report we describe modifications of the original Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) protocol that yields more consistent and less expensive results.

Vídeo: Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY

Multiphoton Microscopy of Cleared Mouse Brain Expressing YFP

Multiphoton microscopy of intrinsic fluorescence and second harmonic generation (SHG) of whole mouse organs is made possible by optically clearing the organ before imaging.1,2 However, for organs that contain fluorescent proteins such as GFP and YFP, optical clearing protocols that use methanol dehydration and clear using benzyl alcohol:benzyl benzoate (BABB) while unprotected from light3 do not preserve the fluorescent signal. The protocol presented here is a novel way in which to perform whole organ optical clearing on mouse brain while preserving the fluorescence signal of YFP expressed in neurons. Altering the optical clearing protocol such that the organ is dehydrated using an ethanol graded series has been found to reduce the damage to the fluorescent proteins and preserve their fluorescent signal for multiphoton imaging.4 Using an optimized method of optical clearing with ethanol-based dehydration and clearing by BABB while shielded from light, we show high-resolution multiphoton images of yellow fluorescent protein (YFP) expression in the neurons of a mouse brain more than 2 mm beneath the tissue surface.

Multiphoton microscopy of whole mouse organs is possible by optically clearing the organ before imaging, but not all protocols preserve the fluorescent signal of fluorescent proteins. Using an optical clearing method with ethanol-based dehydration and benzyl alcohol:benzyl benzoate clearing, we show high-resolution multiphoton images of whole mouse brain expressing YFP.

Microscopia multifotônica do cérebro do rato Cleared Expressando YFP

Multiphoton microscopy of intrinsic fluorescence and second harmonic generation (SHG) of whole mouse organs is made possible by optically clearing the ...

Neurociência

A Laser-induced Mouse Model of Chronic Ocular Hypertension to Characterize Visual Defects

Glaucoma, frequently associated with elevated intraocular pressure (IOP), is one of the leading causes of blindness. We sought to establish a mouse model of ocular hypertension to mimic human high-tension glaucoma. Here laser illumination is applied to the corneal limbus to photocoagulate the aqueous outflow, inducing angle closure. The changes of IOP are monitored using a rebound tonometer before and after the laser treatment. An optomotor behavioral test is used to measure corresponding changes in visual capacity. The representative result from one mouse which developed sustained IOP elevation after laser illumination is shown. A decreased visual acuity and contrast sensitivity is observed in this ocular hypertensive mouse. Together, our study introduces a valuable model system to investigate neuronal degeneration and the underlying molecular mechanisms in glaucomatous mice.

Chronic ocular hypertension is induced using laser photocoagulation of the trabecular meshwork in mouse eyes. The intraocular pressure (IOP) is elevated for several months after laser treatment. The decrease of visual acuity and contrast sensitivity of experimental animals are monitored using the optomotor test.

Um modelo de rato induzida por laser de crônica Ocular Hypertension para Caracterizar defeitos visuais

Glaucoma, frequently associated with elevated intraocular pressure (IOP), is one of the leading causes of blindness. We sought to establish a mouse model ...

Medicina

Micron-scale Resolution Optical Tomography of Entire Mouse Brains with Confocal Light Sheet Microscopy

Understanding the architecture of mammalian brain at single-cell resolution is one of the key issues of neuroscience. However, mapping neuronal soma and projections throughout the whole brain is still challenging for imaging and data management technologies. Indeed, macroscopic volumes need to be reconstructed with high resolution and contrast in a reasonable time, producing datasets in the TeraByte range. We recently demonstrated an optical method (confocal light sheet microscopy, CLSM) capable of obtaining micron-scale reconstruction of entire mouse brains labeled with enhanced green fluorescent protein (EGFP). Combining light sheet illumination and confocal detection, CLSM allows deep imaging inside macroscopic cleared specimens with high contrast and speed. Here we describe the complete experimental pipeline to obtain comprehensive and human-readable images of entire mouse brains labeled with fluorescent proteins. The clearing and the mounting procedures are described, together with the steps to perform an optical tomography on its whole volume by acquiring many parallel adjacent stacks. We showed the usage of open-source custom-made software tools enabling stitching of the multiple stacks and multi-resolution data navigation. Finally, we illustrated some example of brain maps: the cerebellum from an L7-GFP transgenic mouse, in which all Purkinje cells are selectively labeled, and the whole brain from a thy1-GFP-M mouse, characterized by a random sparse neuronal labeling.

In this article we describe the full experimental procedure to reconstruct, with high resolution, the fine brain anatomy of fluorescently labeled mouse brains. The described protocol includes sample preparation and clearing, specimen mounting for imaging, data post-processing and multi-scale visualization.

Micron escala Resolução óptica Tomografia de todo cérebro do rato com confocal Luz Folha de Microscopia

Understanding the architecture of mammalian  brain at single-cell resolution is one of the key issues of neuroscience.  However, mapping neuronal soma and ...

Imaging Serotonergic Fibers in the Mouse Spinal Cord Using the CLARITY/CUBIC Technique

Long descending fibers to the spinal cord are essential for locomotion, pain perception, and other behaviors. The fiber termination pattern in the spinal cord of the majority of these fiber systems have not been thoroughly investigated in any species. Serotonergic fibers, which project to the spinal cord, have been studied in rats and opossums on histological sections and their functional significance has been deduced based on their fiber termination pattern in the spinal cord. With the development of CLARITY and CUBIC techniques, it is possible to investigate this fiber system and its distribution in the spinal cord, which is likely to reveal previously unknown features of serotonergic supraspinal pathways. Here, we provide a detailed protocol for imaging the serotonergic fibers in the mouse spinal cord using the combined CLARITY and CUBIC techniques. The method involves perfusion of a mouse with a hydrogel solution and clarification of the tissue with a combination of clearing reagents. Spinal cord tissue was cleared in just under two weeks, and the subsequent immunofluorescent staining against serotonin was completed in less than ten days. With a multi-photon fluorescent microscope, the tissue was scanned and a 3D image was reconstructed using Osirix software.

Supraspinal projections are important for pain perception and other behaviors, and serotonergic fibers are one of these fiber systems. The present study focused on the application of the combined CLARITY/CUBIC protocol to the mouse spinal cord in order to investigate the termination of these serotonergic fibers.

Imagiologia Fibras Serotoninérgicos na medula espinhal mouse usando o CLARIDADE / Técnica CUBIC

Long descending fibers to the spinal cord are essential for locomotion, pain perception, and other behaviors. The fiber termination pattern in the spinal ...

Induction of Paralysis and Visual System Injury in Mice by T Cells Specific for Neuromyelitis Optica Autoantigen Aquaporin-4

While it is recognized that aquaporin-4 (AQP4)-specific T cells and antibodies participate in the pathogenesis of neuromyelitis optica (NMO), a human central nervous system (CNS) autoimmune demyelinating disease, creation of an AQP4-targeted model with both clinical and histologic manifestations of CNS autoimmunity has proven challenging. Immunization of wild-type (WT) mice with AQP4 peptides elicited T cell proliferation, although those T cells could not transfer disease to na&#239;ve recipient mice. Recently, two novel AQP4 T cell epitopes, peptide (p) 135-153 and p201-220, were identified when studying immune responses to AQP4 in AQP4-deficient (AQP4-/-) mice, suggesting T cell reactivity to these epitopes is normally controlled by thymic negative selection. AQP4-/- Th17 polarized T cells primed to either p135-153 or p201-220 induced paralysis in recipient WT mice, that was associated with predominantly leptomeningeal inflammation of the spinal cord and optic nerves. Inflammation surrounding optic nerves and involvement of the inner retinal layers (IRL) were manifested by changes in serial optical coherence tomography (OCT). Here, we illustrate the approaches used to create this new in vivo model of AQP4-targeted CNS autoimmunity (ATCA), which can now be employed to study mechanisms that permit development of pathogenic AQP4-specific T cells and how they may cooperate with B cells in NMO pathogenesis.

Here, we present a protocol to induce paralysis and opticospinal inflammation by transfer of aquaporin-4 (AQP4)-specific T cells from AQP4-/- mice into WT mice. In addition, we demonstrate how to use serial optical coherence tomography to monitor visual system dysfunction.

Indução de paralisia e lesão do sistema Visual em camundongos por específicos de células T para Neuromielite Óptica Autoantigen Aquaporin-4

While it is recognized that aquaporin-4 (AQP4)-specific T cells and antibodies participate in the pathogenesis of neuromyelitis optica (NMO), a human ...

Imunologia e Infecção

Novel Passive Clearing Methods for the Rapid Production of Optical Transparency in Whole CNS Tissue

Since the development of CLARITY, a bioelectrochemical clearing technique that allows for three-dimensional phenotype mapping within transparent tissues, a multitude of novel clearing methodologies including CUBIC (clear, unobstructed brain imaging cocktails and computational analysis), SWITCH (system-wide control of interaction time and kinetics of chemicals), MAP (magnified analysis of the proteome), and PACT (passive clarity technique), have been established to further expand the existing toolkit for the microscopic analysis of biological tissues. The present study aims to improve upon and optimize the original PACT procedure for an array of intact rodent tissues, including the whole central nervous system (CNS), kidneys, spleen, and whole mouse embryos. Termed psPACT (process-separate PACT) and mPACT (modified PACT), these novel techniques provide highly efficacious means of mapping cell circuitry and visualizing subcellular structures in intact normal and pathological tissues. In the following protocol, we provide a detailed, step-by-step outline on how to achieve maximal tissue clearance with minimal invasion of their structural integrity via psPACT and mPACT.

Here, we present two novel methodologies, psPACT and mPACT, for achieving maximal optical transparency and subsequent microscopic analysis of tissue vasculature in the intact rodent whole CNS.

Passivo novo métodos de compensação para a produção rápida de transparência óptica no tecido inteiro do CNS

Since the development of CLARITY, a bioelectrochemical clearing technique that allows for three-dimensional phenotype mapping within transparent tissues, ...

Immunostaining of Whole-Mount Retinas with the CLARITY Tissue Clearing Method

The tissue hydrogel delipidation method (CLARITY), originally developed by the Deisseroth laboratory, has been modified and widely used for immunostaining and imaging of thick brain samples. However, this advanced technology has not yet been used for whole-mount retinas. Although the retina is partially transparent, its thickness of approximately 200 &#181;m (in mice) still limits the penetration of antibodies into the deep tissue as well as reducing light penetration for high-resolution imaging. Here, we adapted the CLARITY method for whole-mount mouse retinas by polymerizing them with an acrylamide monomer to form a nanoporous hydrogel and then clearing them in sodium dodecyl sulfate to minimize protein loss and avoid tissue damage. CLARITY-processed retinas were immunostained with antibodies for retinal neurons, glial cells, and synaptic proteins, mounted in a refractive index matching solution, and imaged. Our data demonstrate that CLARITY&#160;can improve the quality of standard immunohistochemical staining and imaging for retinal neurons and glial cells in whole-mount preparation. For instance, 3D resolution of fine axon-like and dendritic structures of dopaminergic amacrine cells were much improved by CLARITY. Compared to non-processed whole-mount retinas, CLARITY can reveal immunostaining for synaptic proteins such as postsynaptic density protein 95. Our results show that CLARITY renders the retina more optically transparent after the removal of lipids and preserves fine structures of retinal neurons and their proteins, which can be routinely used for obtaining high-resolution imaging of retinal neurons and their subcellular structures in whole-mount preparation.

Here we present a protocol to adapt the CLARITY method of the brain tissues for whole-mount retinas to improve the quality of standard immunohistochemical staining and high-resolution imaging of retinal neurons and their subcellular structures.

Imunostaining de Retinas de Monte Inteiro com o Método de Limpeza tecidual CLARITY

The tissue hydrogel delipidation method (CLARITY), originally developed by the Deisseroth laboratory, has been modified and widely used for immunostaining ...

Imaging and Quantification of Intact Neuronal Dendrites via CLARITY Tissue Clearing

Brain activity, the electrochemical signals passed between neurons, is determined by the connectivity patterns of neuronal networks, and from the morphology of processes and substructures within these neurons. As such, much of what is known about brain function has arisen alongside developments in imaging technologies that allow further insight into how neurons are organized and connected in the brain. Improvements in tissue clearing have allowed for high-resolution imaging of thick brain slices, facilitating morphological reconstruction and analyses of neuronal substructures, such as dendritic arbors and spines. In tandem, advances in image processing software provide methods of quickly analyzing large imaging datasets. This work presents a relatively rapid method of processing, visualizing, and analyzing thick slices of labeled neural tissue at high-resolution using CLARITY tissue clearing, confocal microscopy, and image analysis. This protocol will facilitate efforts toward understanding the connectivity patterns and neuronal morphologies that characterize healthy brains, and the changes in these characteristics that arise in diseased brain states.

Neuronal dendritic morphology often underlies function. Indeed, many disease processes that affect the development of neurons manifest with a morphological phenotype. This protocol describes a simple and powerful method for analyzing intact dendritic arbors and their associated spines.

Imagem e quantificação de dendritos neuronais intactos via clareza de tecido

Brain activity, the electrochemical signals passed between neurons, is determined by the connectivity patterns of neuronal networks, and from the ...

A Tissue Clearing Method for Neuronal Imaging from Mesoscopic to Microscopic Scales

A detailed protocol is provided here to visualize neuronal structures from mesoscopic to microscopic levels in brain tissues. Neuronal structures ranging from neural circuits to subcellular neuronal structures are visualized in mouse brain slices optically cleared with ScaleSF. This clearing method is a modified version of ScaleS and is a hydrophilic tissue clearing method for tissue slices that achieves potent clearing capability as well as a high-level of preservation of fluorescence signals and structural integrity. A customizable three dimensional (3D)-printed imaging chamber is designed for reliable mounting of cleared brain tissues. Mouse brains injected with an adeno-associated virus vector carrying enhanced green fluorescent protein gene were fixed with 4% paraformaldehyde and cut into slices of 1-mm thickness with a vibrating tissue slicer. The brain slices were cleared by following the clearing protocol, which include sequential incubations in three solutions, namely, ScaleS0 solution, phosphate buffer saline (&#8211;), and ScaleS4 solution, for a total of 10.5&#8211;14.5 h. The cleared brain slices were mounted on the imaging chamber and embedded in 1.5% agarose gel dissolved in ScaleS4D25(0) solution. The 3D image acquisition of the slices was carried out using a confocal laser scanning microscope equipped with a multi-immersion objective lens of a long working distance. Beginning with mesoscopic neuronal imaging, we succeeded in visualizing fine subcellular neuronal structures, such as dendritic spines and axonal boutons, in the optically cleared brain slices. This protocol would facilitate understanding of neuronal structures from circuit to subcellular component scales.

The protocol provides a detailed method of neuronal imaging in brain slice using a tissue clearing method, ScaleSF. The protocol includes brain tissue preparation, tissue clarification, handling of cleared slices and confocal laser scanning microscopy imaging of neuronal structures from mesoscopic to microscopic levels.

Um método de limpeza de tecidos para imagens neuronais de escalas mesoscópicas a microscópicas

A detailed protocol is provided here to visualize neuronal structures from mesoscopic to microscopic levels in brain tissues. Neuronal structures ranging ...

Application of Optical Coherence Tomography to a Mouse Model of Retinopathy

Optical coherence tomography (OCT) offers a noninvasive method for the diagnosis of retinopathy. The OCT machine can capture retinal crosssectional images from which the retinal thickness can be calculated. Although OCT is widely used in clinical practice, its application in basic research is not as prevalent, especially in small animals such as mice. Because of the small size of their eyeballs, it is challenging to conduct fundus imaging examinations in mice. Therefore, a specialized retinal imaging system is required to accommodate OCT imaging on small animals. This article demonstrates a small-animal-specific system for OCT examination procedures and a detailed method for image analysis. The results of retinal OCT examination of very-low-density lipoprotein receptor (Vldlr) knockout mice and C57BL/6J mice are presented. The OCT images of C57BL/6J mice showed retinal layers, while those of Vldlr knockout mice showed subretinal neovascularization and retinal thinning. In summary, OCT examination could facilitate the noninvasive detection and measurement of retinopathy in mouse models.

Here, we describe an in vivo imaging technique using optical coherence tomography to facilitate the diagnosis and quantitative measurement of retinopathy in mice.

Aplicação da Tomografia de Coerência Óptica a um Modelo de Retinopatia de Rato

Optical coherence tomography (OCT) offers a noninvasive method for the diagnosis of retinopathy. The OCT machine can capture retinal crosssectional images ...