Este trabalho foi financiado pelo Instituto Nacional de Distúrbios Neurológicos e Derrame conceder NS081333 (a CMT), concessão Associação NIRG-10-171721 de Alzheimer e Instituto Nacional de Saúde Mental de concessão MH096702 (a UH), e do Instituto Nacional sobre Envelhecimento financiado pela pesquisa da doença de Alzheimer Centro da Universidade de Columbia concessão piloto AG008702 (para YYJ e JB).

Nota: Para toda a experimentação animal, Institucional e diretrizes nacionais para o cuidado e uso de animais de laboratório foram seguidas. 1. Prepare Instrumentos Cirúrgicos e Soluções para Cirurgia <ol><li> Autoclave todos os instrumentos cirúrgicos de aço inoxidável. </li><li> Preparar etanol a 70% por diluição de 200 proof de etanol absoluto com grau molecular de água desionizada destilada estéril. </li><li> Fixe a agulha G 29 para a seringa Hamilton. Limpe o interior do Hamilton seringa e agulha através da elaboração e da água de ejecção nanopura repetidamente durante 1 min. Repita este procedimento com etanol 70%. <ol><li> Retirar o êmbolo e a agulha da seringa de Hamilton. Ar secar as peças em um fluxo laminar vertical S / N. </li><li> Irradiar a agulha e seringa com luz ultravioleta durante 30 minutos antes da utilização. </li></ol></li><li> Preparar a solução salina através da dissolução de NaCl em água de grau molecular para um peso final a concentração em volume de 0,9%. Esterilização emze a solução filtrando-a através dos poros do filtro 0,2 m. </li></ol> 2. Prepare oligomica Ap 1-42 <ol><li> Monomerize e ressuspender recombinante humana Ap 1-42 em DMSO a 5 mM exactamente como descrito na publicação da Fa &quot;e outros 13. </li><li> Diluir 5 mM Ap 1-42 com estéril (1x) PBS 100 mM na véspera da cirurgia. </li><li> Misture bem a solução por trituração. </li><li> Incubar Ap 1-42 solução durante 12 horas a 4 ° C. Nota: As soluções de controle, tais como a diluição de DMSO em PBS ou mexidos / retrocesso Ap 1-42 deve ser preparado da mesma forma que Ap 1-42. </li></ol> 3. determinar as coordenadas de injeção <ol><li> Use os adultos atlas do cérebro do rato para determinar a exacta ântero-posterior (AP), medial-lateral (ML), e dorso-ventral (DV) coordenadas para a região do cérebro de 14 interesse. Nãote: Para ilustrar a técnica, escolhemos o giro denteado do hipocampo como o alvo com as seguintes coordenadas a partir do bregma: AP, -2,00 mm; ML, ± 1,3 mm; DV, -2,2 mm. O sinal negativo que precede o valor AP indica que ele é de 2,00 mm posterior do bregma. O sinal ± precedendo o valor ML indica a direção esquerda e direita do centro. Por fim, o sinal negativo que precede o DV indica que está em movimento ventral a partir da superfície do cérebro. </li></ol> 4. estereotáxica Setup Moldura <ol><li> Usar uma máscara cirúrgica rosto, jaleco limpo, e luvas cirúrgicas estéreis. </li><li> Limpe instrumento estereotáxico e adaptador de mouse com etanol 70%. </li><li> Deite-campo cirúrgico estéril sobre o balcão. </li><li> Colocar o aparelho estereotáxico com um adaptador do rato no topo da cobertura cirúrgica estéril. </li><li> Limpe a almofada de aquecimento do mouse com etanol 70%. Posicione a almofada de aquecimento sobre a cama quadro estereotáxico. Use labora uso geralfita rotulagem tory a fita para baixo a almofada de aquecimento sobre a cama adaptador de mouse. <ol><li> Fixe a almofada de aquecimento para a bomba de água morna recirculator acordo com as instruções do fabricante. </li><li> Ligue a bomba recirculator água morna e definir a temperatura a 37 ° C, e depois esperar até que ele atinja a temperatura. </li></ol></li><li> Apor o 50 ul seringa Hamilton com uma agulha G 29 sobre a armação estereotáxica aparafusando-o sobre o eixo vertical, motorizada de injecção de acordo com as instruções do fabricante. </li><li> Ligue o esterilizador talão e esperar até atingir a temperatura programada pelo fabricante. Nota: Este é usado para esterilizar instrumentos de aço inoxidável entre os animais. Leva instrumentos de 20 segundos de contacto com os grânulos de esterilização. </li><li> Ligue placa quente e configurá-lo para 42 ° C e coloque uma gaiola vazia limpo no topo. </li><li> Enquanto a seringa Hamilton está fixada no quadro estereotáxico usar o i motorizado estereotáxiconjector para elaborar o 1-42 solução de Ap. Nota: Elaborar mais de 4 mL da solução para a seringa e, em seguida, usar o injector estereotáxico para definir o volume de injecção. Operar o injector de acordo com as instruções do fabricante. </li></ol> 5. Preparação de animais <ol><li> Determinar o peso do rato utilizando a escala de pesagem. </li><li> Anestesiar rato com cetamina / xilazina coquetel a 100 mg / kg de cetamina e 10 mg / kg de xilazina por injecção intraperitoneal (IP). Monitore a profundidade da anestesia com a perda de toe pitada reflexo. </li><li> Após o mouse está sedado tomar a máquina de cortar cabelo e fazer a barba a cabeça para expor a pele ao longo do crânio. </li><li> Colocar o rato no topo da almofada de aquecimento na parte superior do leito estereotáxico. </li><li> Use a espátula para abrir a boca e colocar os dentes incisivos dentro do guarda-dentes. Prenda o porta-objetivas sobre a face do mouse. Apertá-lo com cuidado e não muito apertado. </li><li> Posição as travessas de ouvido bilaterais em conduto auditivo para proteger a cabeça. Mova cada barra transversal até que ela atinge o crânio, e, em seguida, gire o parafuso para prendê-la. <ol><li> Para garantir que a cabeça está garantido usar o dedo indicador para empurrar suavemente para baixo na cabeça. Se a cabeça está devidamente assegurado que não vai dar para fora ou mover quando pressionado. </li></ol></li><li> Aplique uma gota de creme para os olhos ou colírio para os olhos para mantê-los úmidos. Certifique-se de que os olhos são úmido durante todo o procedimento cirúrgico. </li><li> Para desinfectar o local cirúrgico usar cotonetes estéreis para aplicar a solução de betadine para a pele sobre o crânio, seguido de etanol a 70%. Realize o betadine alternada e 70% de etanol limpeza mais 2 vezes. </li></ol> 6. Cirurgia e Infusão <ol><li> Use um bisturi para fazer uma incisão de 2-3 mm na linha média do couro cabeludo, e então usar uma tesoura fina reta para estender a linha de incisão para 1,0-1,5 cm para expor a sutura sagital, bregma e lambda do crânio, marcos as coordenadas estereotáxicas que se baseiam. Use micro grampos para manter a pele separados. Nota: As posições bregma e lambda são explicados no atlas do cérebro do rato &quot;O cérebro do rato em estereotáxica coordenadas&quot; 14. </li><li> Pegue um cotonete, mergulhe-o em solução salina estéril, e usá-lo para limpar o crânio. Em seguida, use um cotonete seco e limpo para secar o crânio. </li><li> Use uma caneta de ponta fina estéril para marcar um ponto no bregma. Utilizar o micromanipulador estereotáxico para posicionar a seringa de Hamilton para que a ponta da agulha apenas toca o ponto no bregma. <ol><li> Grave coordenar a DV. </li><li> Para verificar se o eixo PA do nível do crânio é mover a seringa Hamilton posteriormente de modo a que a ponta da agulha atinge o ponto de lambda. </li><li> Grave a posição DV. Certifique-se coordena o DV para bregma e lambda são de 0,5 mm. Nota: O objectivo é minimizar a diferença entre DV bregma e lambda. </li></ol></li><li> Usa-osicromanipulator para retornar a agulha da seringa Hamilton ponta de volta para o ponto bregma. </li><li> Grave a posição inicial AP. </li><li> Calcular a posição final AP subtraindo 2,00 milímetros a partir da AP a partir de coordenadas. </li><li> Use o micromanipulador para reposicionar a agulha de seringa Hamilton com a AP cota final. </li><li> Grave coordenar o atual ML. </li><li> Calcule a esquerda e direita que termina ML coordena adicionando ou subtraindo 1,3 milímetros a partir da partida ML coordenadas. </li><li> Use o micromanipulador para mover a agulha da seringa Hamilton, quer coordenar a ML terminando. <ol><li> Toque a ponta da agulha para a superfície do crânio em ambos terminam coordenadas ML e gravar as coordenadas DV e certifique-se os valores estão dentro de 0,5 mm. Nota: O objectivo é minimizar a diferença entre os dois DV terminando coordenadas ML. </li><li> Enquanto em um final de coordenadas puxar a agulha para cima 0,5-1 cm sobre o crânio. Pegue uma caneta de ponta fina estéril para marcar um ponto no the a superfície do crânio. Repita este passo para o lado contralateral do crânio. </li></ol></li><li> Balançar a seringa Hamilton clara para fora do caminho. </li><li> Anexar 0,8 milímetros cabeça de perfuração para a broca. Pegue a furadeira com ambas as mãos, defina os cotovelos sobre a superfície da mesa para a estabilidade, e posicionar a cabeça da broca ligeiramente acima do ponto sharpie sobre a esquerda ou direita ML coordenadas. Activa a broca, diminuir a ponta da broca para a superfície do crânio para introduzir um orifício no crânio. Repita este passo para o lado contralateral. Nota: Alguns hemorragia pode ocorrer a partir dos furos recentemente introduzidos. Se houver sangramento, em seguida, use um cotonete limpo e seco para enxugar o sangue. </li><li> Mova a seringa Hamilton de volta para a posição sobre um dos buracos ML recentemente introduzidas. Abaixe a agulha Hamilton apenas após o crânio. Neste ponto, deve ter cuidado para não perfurar o cérebro. Para certificar-se de que a agulha passou o crânio levar o seu dedo indicador e empurre a agulha contra o crânio para fazer sure a agulha não sair do furo. </li><li> Grave coordenar o DV partida. </li><li> Calcule coordenar o DV terminando subtraindo 2,2 milímetros do DV começando coordenadas. </li><li> Abaixe a agulha até o DV terminando coordenadas. </li><li> Activar a bomba injectora estereotáxico para bombear 4 ul de Ap 1-42 no giro dentado, a uma taxa de 0,5 mL / min. </li><li> Quando a conclusão da perfusão deixar a agulha permanecer no lugar durante um 1 minuto adicional de minimizar o refluxo da solução para fora do local de injecção. </li><li> Mover a agulha para o outro lado do cérebro e repetir as etapas de 6,13-6,18. </li></ol> 7. Remoção Animal e Cuidados Pós-Operatórios <ol><li> Soltar os bares de ouvido e proteção para o rosto / nariz. Retirar o mouse do aparelho. </li><li> Use um estudante fórceps padrão normal para puxar fechar o couro cabeludo e, em seguida, selar a ferida com sutura. </li><li> Injectar 1 ml de solução salina estéril para o mouse via IP para a hidratação. </li><li> Emject buprenorfina (0,1 mg / kg) por via subcutânea para aliviar a dor. </li><li> Manter o rato na gaiola vazia limpo situado no topo de 42 ° C chapa quente até que ele acorda, em seguida, devolvê-lo à sua gaiola de habitação com amplo comida e água. </li><li> Administrar buprenorfina, por injecção subcutânea a cada 12 horas durante 3 dias para alívio da dor. </li></ol> Remoção 8. Sutura <ol><li> Anestesiar rato com cetamina / xilazina coquetel a 100 mg / kg de cetamina e 10 mg / kg xlyazine por IP. Nota: Isto é feito 7-10 dias após a cirurgia. </li><li> Use um estudante fórceps padrão standard e uma tesoura fina reta para remover a sutura. </li><li> Injectar 1 ml de solução salina estéril para o mouse via IP para a hidratação. </li><li> Manter o rato em uma gaiola vazia limpo situado no topo de 42 ° C chapa quente até que ele acorda, e, em seguida, devolvê-lo à sua gaiola de habitação com amplo comida e água. </li></ol> 9. sacrifício de animais e processamento de tecidos <ol><li> Anestesiar o mouse e perfuse com paraformaldeído a 4% 15, remover o cérebro, e processá-lo para cryosectioning 16. Nota: O tempo de sacrifício de animais depende do experimentador. No entanto, para os nossos estudos, escolheu 7 e 14 dias após a cirurgia. </li></ol>

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R., Zaborszky, L., Heimer, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distribution+of+GABAergic+and+cholinergic+neurons+in+the+rat+diagonal+band.">Distribution of GABAergic and cholinergic neurons in the rat diagonal band.</a> Neuroscience. 17 (2), 439-451 (1986).</li><li>Clarke, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cholinergic+innervation+of+the+rat+dentate+gyrus:+an+immunocytochemical+and+electron+microscopical+study.">Cholinergic innervation of the rat dentate gyrus: an immunocytochemical and electron microscopical study.</a> Brain research. 360 (1-2), 349-354 (1985).</li><li>Garcia-Osta, A., Alberini, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Amyloid+beta+mediates+memory+formation.">Amyloid beta mediates memory formation.</a> Learning &amp; memory. 16 (4), 267-272 (2009).</li></ol>

Os autores declaram que não têm interesses financeiros concorrentes.

Para conseguir um sucesso Ap 1-42 injeção do experimentador ou cirurgião deve: 1) utilizar uma técnica asséptica; 2) identificar corretamente a região do cérebro de interesse com coordenadas precisas; 3) ser capaz de garantir adequadamente o mouse no quadro estereotáxico com o cérebro nivelado no eixo AP e ML; 4) tem a capacidade de operar o micromanipulador com precisão; 5) Garantir cuidados pós-operatórios adequada. Se essas etapas principais são seguidos do mouse deve sobreviver à cirurgia se...

Depósitos de placas amilóides, que são compostos de beta-amilóide (Ap 1-42), são um elemento-chave da patologia da doença de Alzheimer (DA). Numerosos estudos têm demonstrado que níveis elevados ou tóxicos de recombinante oligomeric Ap 1-42 provocar a morte neuronal, sináptica distrofia, perda e disfunção; bem como de aprendizagem e memória défices 1-4. Regiões do cérebro afetadas incluem o hipocampo, no córtex e estruturas subcorticais tais como o cérebro anterior basal ea 5,6 amígdala. Até à data, existem vários modelos de camundongos transgênicos que tentam simular o Aâ 1-42 patologia da AD. Dependendo da estirpe estes animais revelar-se útil para examinar seleccione características patológicas da DA. Infelizmente, com a excepção de duas linhas transgénicas, APP23 e 5XFAD, estes ratinhos não replicar completamente a perda neuronal, um aspecto chave da AD. Mesmo com a perda neuronal observada em APP23 e 5XFAD, a obser morte neuronalved foi sutil, dependente da idade, e isolado para algumas regiões selecione 7,8. A infusão direta de oligomeric Ap 1-42 no cérebro do rato do tipo selvagem é um excelente modelo in vivo que replica o aspecto morte neuronal de amyloidopathy 1,9,10. Ao contrário dos modelos de ratinho transgénicos vulgarmente utilizados o modelo de infusão oligomérico Ap 1-42 é ideal para elevar de forma aguda Ap 1-42 níveis de um modo espacial e temporal. A vantagem da utilização de ratinhos de tipo selvagem para este modelo de compensação ou elimina efeitos colaterais potenciais de mutações introduzidas em linhas de ratos transgénicos. Estudos anteriores têm demonstrado que a infusão de níveis tóxicos de Ap 1-42 no hipocampo provoca a morte neuronal na proximidade do local de injecção dentro de 1 semana 1. Além disso, consistente com a observação de que Ap 1-42 é tóxico para os neurónios colinérgicos do prosencéfalo basal 11população colinérgicos neurônio (BFCN) que se projecta para o hipocampo diminui 20-50% no prazo de 7-14 dias após a beta-amilóide infusão 1,10 em camundongos, de forma eficaz permitindo que para os exames de circuitos neuronais isolado no cérebro. Desde projecto BFCN ipsilateralmente ao giro dentado do hipocampo 12, para a maior parte de controlo / veículo e oligoméricos Ap 1-42 soluções podem ser injectados em ambos os lados do cérebro, permitindo que sejam feitas comparações entre os hemisférios direito e esquerdo 1. Neste relatório, iremos fornecer uma metodologia cirúrgica ea injeção detalhado para adultos de tipo selvagem camundongos C57BL / 6J. Esta estirpe do mouse é escolhido por causa de sua ampla utilização na pesquisa. Tecnicamente, qualquer região do cérebro podem ser direcionados para perfusão, no entanto aqui, vamos utilizar o giro denteado do hipocampo como o alvo para ilustrar a técnica.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Ketamine HCl (100mg/ml)</td>
			<td>Henry Schein Medical</td>
			<td>1049007</td>
			<td>100 mg ketamine per 1 kg animal</td>
		</tr>
		<tr>
			<td>Xylazine (20mg/ml)</td>
			<td>Henry Schein Medical</td>
			<td>not available</td>
			<td>10 mg xylazine per 1 kg animal</td>
		</tr>
		<tr>
			<td>Buprenex (0.3mg/ml)</td>
			<td>Henry Schein Medical</td>
			<td>1217793</td>
			<td>0.1 mg buprenex per 1 kg animal</td>
		</tr>
		<tr>
			<td>A&#946;1-42</td>
			<td>David Teplow/UCLA</td>
			<td>not available</td>
			<td>100 &#956;M; This amyloid was used in the paper</td>
		</tr>
		<tr>
			<td>A&#946;1-42</td>
			<td>Bachem</td>
			<td>H-1368</td>
			<td>Can be used in place of amyloid from the Teplow lab</td>
		</tr>
		<tr>
			<td>A&#946;1-42</td>
			<td>American Peptide</td>
			<td>62-0-80B</td>
			<td>Can be used in place of amyloid from the Teplow lab</td>
		</tr>
		<tr>
			<td>Scrambled A&#946;1-42</td>
			<td>American Peptide</td>
			<td>62-0-46B</td>
			<td>Can be used as control peptide for comparing A&#946;1-42</td>
		</tr>
		<tr>
			<td>NU4 Antibody (Oligomeric Amyloid Antibody)</td>
			<td>Gift from William Klein/Northwestern U.</td>
			<td>not available</td>
			<td>1:2000 dilution</td>
		</tr>
		<tr>
			<td>Anti-Amyloid Oligomeric Antibody&#160; (Polyclonal Rabbit)</td>
			<td>EMD Millipore</td>
			<td>AB9234</td>
			<td>May be used in place of Nu4; needs to&#160; be tested by the end user</td>
		</tr>
		<tr>
			<td>6E10 Antibody (Monoclonal Mouse) (Amyloid Antibody)</td>
			<td>Biolegend</td>
			<td>sig-39320</td>
			<td>1:1000 dilution</td>
		</tr>
		<tr>
			<td>ChAT Antibody (Polyclonal Goat)</td>
			<td>Millipore</td>
			<td>AB144P</td>
			<td>1:100 dilution</td>
		</tr>
		<tr>
			<td>DeadEnd Fluorometric TUNEL system</td>
			<td>Promega</td>
			<td>G3250</td>
			<td>Follow manufacturer&#39;s directions for use</td>
		</tr>
		<tr>
			<td>Prolong Gold Antifade Reagent with DAPI</td>
			<td>Invitrogen</td>
			<td>P36935</td>
			<td>Use when coverslipping slides</td>
		</tr>
		<tr>
			<td>Fluorogold</td>
			<td>Fluorochrome, LLC</td>
			<td>not available</td>
			<td>2% solution</td>
		</tr>
		<tr>
			<td>Absolute Ethanol (200 proof)</td>
			<td>Fisher Scientific</td>
			<td>BP2818-4</td>
			<td>For making 70% ethanol for sanitizing and disinfecting</td>
		</tr>
		<tr>
			<td>Novex 10-20% Tricine gel</td>
			<td>Life Technologies</td>
			<td>EC6625BOX</td>
			<td>For separating A&#946;1-42</td>
		</tr>
		<tr>
			<td>Novex Tricine SDS Running Buffer (10X)</td>
			<td>Life Technologies</td>
			<td>LC1675</td>
			<td>For running 10-20% Tricine gels</td>
		</tr>
		<tr>
			<td>Novex Tris-Glycine Transfer Buffer (25X)</td>
			<td>Life Technologies</td>
			<td>LC3675</td>
			<td>For transferring 10-20% Tricine gels</td>
		</tr>
		<tr>
			<td>SuperSignal Western Blot Enhancer</td>
			<td>Thermo Scientific</td>
			<td>46640</td>
			<td>For enhancing A&#946;1-42 signal; follow manufacturer&#39;s protocol</td>
		</tr>
		<tr>
			<td>Protran BA79 Nitrocellulose Blotting Membrane, 0.1 &#956;m</td>
			<td>GE Healthcare Life Sciences</td>
			<td>10402088</td>
			<td>For transferring 10-20% Tricine gels</td>
		</tr>
		<tr>
			<td>Xcell SureLock Mini-Cell</td>
			<td>Life Technologies</td>
			<td>EI0001</td>
			<td>Electrophoresis aparatus for running 10-20% Tricine gels</td>
		</tr>
		<tr>
			<td>GenTeal Lubricant Eye Gel</td>
			<td>Novartis</td>
			<td>not available</td>
			<td>For keeping the mouse eyes moist during surgery; can be found in local pharmacy stores</td>
		</tr>
		<tr>
			<td>Refresh Optive Lubricant Eye Drops</td>
			<td>Allergan</td>
			<td>not available</td>
			<td>For keeping the mouse eyes moist during surgery; can be found in local pharmacy stores; Can be used in place of GenTeal</td>
		</tr>
		<tr>
			<td>Betadine</td>
			<td>Stoelting</td>
			<td>50998</td>
			<td>For sanitizing and disinfecting</td>
		</tr>
		<tr>
			<td>Round/Tapered Spatula&#160;</td>
			<td>VWR</td>
			<td>82027-490</td>
			<td>For opening animal mouth</td>
		</tr>
		<tr>
			<td>Bulldog Serrefines Clamps (Jaw Dims. 9X1.6mm; Length 28mm)</td>
			<td>Fine Science Tools</td>
			<td>18050-28</td>
			<td>Optional; For keeping scalp skin apart during injection</td>
		</tr>
		<tr>
			<td>Straight Fine Scissors (Cutting edge 25mm; Length 11.5cm)</td>
			<td>Fine Science Tools</td>
			<td>14060-11</td>
			<td>For cutting scalp</td>
		</tr>
		<tr>
			<td>#3 Scalpel Handle</td>
			<td>Fine Science Tools</td>
			<td>10003-12</td>
			<td></td>
		</tr>
		<tr>
			<td>#11 Surgical Blade</td>
			<td>Fine Science Tools</td>
			<td>10011-00</td>
			<td>For making scalp incision</td>
		</tr>
		<tr>
			<td>Student Standard Pattern Forcep (Tip Dims. 2.5x1.5mm; Length 11.5cm)</td>
			<td>Fine Science Tools</td>
			<td>91100-12</td>
			<td>For holding scalp closed during suturing</td>
		</tr>
		<tr>
			<td>Trimmer Combo Kit</td>
			<td>Kent Scientific</td>
			<td>CL9990-1201</td>
			<td>For shaving hair</td>
		</tr>
		<tr>
			<td>T/Pump Warm Water Recirculator&#160;</td>
			<td>Kent Scientific&#160;</td>
			<td>TP-700</td>
			<td>For warming animal during surgery</td>
		</tr>
		<tr>
			<td>Resusable Warmining Pad (5&#34; x 10&#34;)</td>
			<td>Kent Scientific&#160;</td>
			<td>TPZ-0510FEA</td>
			<td>For attaching it to the T/Pump warm water recirculator to warm the animal during surgery</td>
		</tr>
		<tr>
			<td>Cordless Micro Drill</td>
			<td>Stoelting</td>
			<td>58610</td>
			<td>Use 0.8mm steel burrs to drill holes in the skull</td>
		</tr>
		<tr>
			<td>Lab Standard Stereotaxic Instrument with Mouse &#38; Neonatal Rat Adaptor</td>
			<td>Stoelting</td>
			<td>51615</td>
			<td></td>
		</tr>
		<tr>
			<td>Just for Mouse Stereotaxic Instrument</td>
			<td>Stoelting</td>
			<td>51730</td>
			<td>Can use this in place of Stoelting Cat. #51615</td>
		</tr>
		<tr>
			<td>Quintessential Stereotaxic Injector</td>
			<td>Stoelting</td>
			<td>53311</td>
			<td></td>
		</tr>
		<tr>
			<td>Dry Glass Bead Sterilizer</td>
			<td>Stoelting</td>
			<td>50287</td>
			<td>For sterilizing stainless steel instruments</td>
		</tr>
		<tr>
			<td>Sterile Surgical Drape (18&#34; x 26&#34;)</td>
			<td>Stoelting</td>
			<td>50981</td>
			<td></td>
		</tr>
		<tr>
			<td>Hamilton Syringe 50 ml, Model 705 RN SYR, NDL</td>
			<td>Hamilton Company</td>
			<td>7637-01</td>
			<td>For brain injection; use different syringes for different solutions</td>
		</tr>
		<tr>
			<td>29 Gauge Needle, Small Hub RN NDL</td>
			<td>Hamilton Company</td>
			<td>7803-06</td>
			<td>For attaching to the Hamilton syringe for brain injection</td>
		</tr>
		<tr>
			<td>1 ml BD Tuberculin Syringes</td>
			<td>VWR</td>
			<td>BD309659</td>
			<td>For administering anesthesia and saline</td>
		</tr>
		<tr>
			<td>30 Gauge Needle (0.5&#34;)</td>
			<td>VWR</td>
			<td>BD305106</td>
			<td>For administering anesthesia and saline</td>
		</tr>
		<tr>
			<td>Portable Electronic CS Series Scale (Ohaus)</td>
			<td>VWR</td>
			<td>65500-202</td>
			<td>For weighing animals to determine anesthesia dose</td>
		</tr>
		<tr>
			<td>Hot plate (Top Plate Dims. 7.25x7.25in)</td>
			<td>VWR</td>
			<td>47751-148</td>
			<td>For warming animals post-surgery</td>
		</tr>
		<tr>
			<td>Sofsilk Silk Suture C-1 Cutting 3/8, 12 mm</td>
			<td>Covidien</td>
			<td>S1173</td>
			<td>For closing wound</td>
		</tr>
		<tr>
			<td>Vetbond Tissue Adhesive (3M)</td>
			<td>Santa Cruz Biotechnology</td>
			<td>sc-361931</td>
			<td>Optional: for aiding in wound closure; Use with suture.</td>
		</tr>
		<tr>
			<td>Cotton-Tipped Wooden-Shaft Sterile Applicators</td>
			<td>Fisher scientific</td>
			<td>22-029-488</td>
			<td>For cleaning and drying surgical wound</td>
		</tr>
		<tr>
			<td>Fisherbrand Superfrost Plus Microscope Slides</td>
			<td>Fisher Scientific&#160;</td>
			<td>12-550-15</td>
			<td>For collecting brain sections</td>
		</tr>
		<tr>
			<td>VWR Micro Cover Glass 24 X 50 mm</td>
			<td>VWR</td>
			<td>48393241</td>
			<td>For mounting microscope slides</td>
		</tr>
		<tr>
			<td>Thermo Scientific Nalgene Syringe Filter 0.2 &#956;m</td>
			<td>Fisher Scientific</td>
			<td>194-2520</td>
			<td>For sterilizing saline solution</td>
		</tr>
		<tr>
			<td>Sterile dual tip skin markers by Viscot Medical</td>
			<td>Medline</td>
			<td>VIS1422SRL91</td>
			<td>For marking coordinates on the skull</td>
		</tr>
	</tbody>
</table>

stereotaxic infusion of oligomeric amyloid-beta into the mouse hippocampus

A doença de Alzheimer é uma doença neurodegenerativa que afecta o envelhecimento da população. Uma característica neuropatológica chave da doença é a sobre-produção de beta-amilóide e a deposição de placas de amilóide-beta em regiões do cérebro dos indivíduos afectados. Ao longo dos anos, os cientistas têm gerado numerosos modelos de rato da doença de Alzheimer que tentam reproduzir a patologia beta-amilóide. Infelizmente, os modelos de ratinho única imitar selectivamente as características da doença. A morte neuronal, um efeito proeminente nos cérebros de pacientes com doença de Alzheimer, é notoriamente ausente nestes ratinhos. Por isso, nós e outros têm utilizado um método de infundir diretamente espécies oligoméricas solúveis de beta-amilóide - Formas de amilóide-beta que têm sido provada a ser mais tóxico para os neurônios - stereotaxically no cérebro. Neste relatório nós utilizamos camundongos C57BL / 6J para documentar esta técnica cirúrgica de aumentar os níveis de beta-amilóide em uma região do cérebro select. Oinfusão alvo é o giro dentado do hipocampo, porque esta estrutura do cérebro, juntamente com o cérebro anterior basal, que é ligado pelo circuito colinérgico, representa uma das áreas de degeneração na doença. Os resultados de elevação beta-amilóide no giro denteado através de infusão estereotáxico revelam aumentos na perda de neurónios no giro dentado no prazo de 1 semana, enquanto existe um aumento concomitante na morte de células e perda de neurónios colinérgicos no membro vertical da banda diagonal de Broca do prosencéfalo basal. Estes efeitos são observados até 2 semanas. Nossos dados sugerem que o atual modelo de infusão de beta-amilóide fornece um modelo alternativo do mouse para abordar a morte dos neurônios específicos região em uma base de curto prazo. A vantagem deste modelo é que a beta-amilóide pode ser elevada de um modo espacial e temporal.

O presente método de preparação de humano recombinante oligomérico Ap 1-42 produz espécies oligoméricas solúveis constituídos por monómeros, dímeros, trímeros e tetrâmeros (Figura 1A). Estes baixo peso molecular de espécies Aâ 1-42, mas não as fibrilas e placas, foram mostrados em várias configurações para ser mais tóxico para os neurônios 1,4,9,17-19. Para determinar se ou não oligomeric Ap 1-42 induz a morte dos neurônios no cérebro do...

Watch this Scientific Journal Video about Stereotaxic Infusion of Oligomeric Amyloid-beta into the Mouse Hippocampus at JoVE.com

Stereotaxic Infusion of Oligomeric Amyloid-beta into the Mouse Hippocampus

Here, we present a protocol for direct stereotaxic brain infusion of amyloid-beta. This methodology provides an alternative in vivo mouse model to address the short-term effects of amyloid-beta on brain neurons.

Infusão estereotáxica de oligomica beta-amilóide no ratinho Hippocampus

Alzheimer&#8217;s disease is a neurodegenerative disease affecting the aging population. A key neuropathological feature of the disease is the ...

stereotaxic-infusion-oligomeric-amyloid-beta-into-mouse

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Neuroscience

18.0K visualizações.  Columbia University Medical Center. Here, we present a protocol for direct stereotaxic brain infusion of amyloid-beta. This methodology provides an alternative in vivo mouse model to address the short-term effects of amyloid-beta on brain neurons.

Vídeo: Stereotaxic Infusion of Oligomeric Amyloid-beta into the Mouse Hippocampus

Ultrasound-Guided Microinjection into the Mouse Forebrain In Utero at E9.5

In utero survival surgery in mice permits the molecular manipulation of gene expression during development. However, because the uterine wall is opaque during early embryogenesis, the ability to target specific parts of the embryo for microinjection is greatly limited. Fortunately, high-frequency ultrasound imaging permits the generation of images that can be used in real time to guide a microinjection needle into the embryonic region of interest. Here we describe the use of such imaging to guide the injection of retroviral vectors into the ventricular system of the mouse forebrain at embryonic day (E) 9.5. This method uses a laparotomy to permit access to the uterine horns, and a specially designed plate that permits host embryos to be bathed in saline while they are imaged and injected. Successful surgeries often result in most or all of the injected embryos surviving to any subsequent time point of interest (embryonically or postnatally). The principles described here can be used with slight modifications to perform injections into the amnionic fluid of E8.5 embryos (thereby permitting infection along the anterior posterior extent of the neural tube, which has not yet closed), or into the ventricular system of the brain at E10.5/11.5. Furthermore, at mid-neurogenic ages (~E13.5), ultrasound imaging can be used direct injection into specific brain regions for viral infection or cell transplantation. The use of ultrasound imaging to guide in utero injections in mice is a very powerful technique that permits the molecular and cellular manipulation of mouse embryos in ways that would otherwise be exceptionally difficult if not impossible.

Ultrasound-Guided Microinjection into the Mouse Forebrain In Utero at E9.5

In utero survival surgery in mice permits the molecular manipulation of gene expression during development. Here we describe the use of high-frequency ultrasound imaging to guide the injection of retroviral vectors into the mouse brain at embryonic day (E) 9.5.

Ultra-som guiada microinjeção em parte frontal do cérebro do rato In Utero Em E9.5

In utero survival surgery in mice permits the molecular manipulation of gene expression during development.  However, because the uterine wall is opaque ...

Neurociência

Stereotaxic Surgery for Excitotoxic Lesion of Specific Brain Areas in the Adult Rat

Many behavioral functions in mammals, including rodents and humans, are mediated principally by discrete brain regions. A common method for discerning the function of various brain regions for behavior or other experimental outcomes is to implement a localized ablation of function. In humans, patient populations with localized brain lesions are often studied for deficits, in hopes of revealing the underlying function of the damaged area. In rodents, one can experimentally induce lesions of specific brain regions.
Lesion can be accomplished in several ways. Electrolytic lesions can cause localized damage but will damage a variety of cell types as well as traversing fibers from other brain regions that happen to be near the lesion site. Inducible genetic techniques using cell-type specific promoters may also enable site-specific targeting. These techniques are complex and not always practical depending on the target brain area. Excitotoxic lesion using stereotaxic surgery, by contrast, is one of the most reliable and practical methods of lesioning excitatory neurons without damaging local glial cells or traversing fibers.
Here, we present a protocol for stereotaxic infusion of the excitotoxin, N-methyl-D-aspartate (NMDA), into the basolateral amygdala complex. Using anatomical indications, we apply stereotaxic coordinates to determine the location of our target brain region and lower an injection needle in place just above the target. We then infuse our excitotoxin into the brain, resulting in excitotoxic death of nearby neurons. While our experimental subject of choice is a rat, the same methods can be applied to other mammals, with the appropriate adjustments in equipment and coordinates.
This method can be used on a variety of brain regions, including the basolateral amygdala1-6, other amygdala nuclei6, 7, hippocampus8, entorhinal cortex9 and prefrontal cortex10. It can also be used to infuse biological compounds such as viral vectors1, 11. The basic stereotaxic technique could also be adapted for implantation of more permanent osmotic pumps, allowing more prolonged exposure to a compound of interest.

Targeted ablation of specific brain region(s) by infusion of an excitotoxin using stereotaxic coordinates is described. This technique could also be adapted for infusion of other chemicals into the rat brain.

Cirurgia estereotáxica para lesão excitotóxica de áreas específicas do cérebro do rato adulto

Many behavioral functions in mammals, including rodents and humans, are mediated principally by discrete brain regions. A common method for discerning the ...

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date this tool has been widely used to study the regulation of cellular proliferation, differentiation and neuronal migration especially in the developing cerebral cortex 6-8. Here we detail our protocol to electroporate in utero the cerebral cortex and the hippocampus and provide evidence that this approach can be used to study dendrites and spines in these two cerebral regions.
Visualization and manipulation of neurons in primary cultures have contributed to a better understanding of the processes involved in dendrite, spine and synapse development. However neurons growing in vitro are not exposed to all the physiological cues that can affect dendrite and/or spine formation and maintenance during normal development. Our knowledge of dendrite and spine structures in vivo in wild-type or mutant mice comes mostly from observations using the Golgi-Cox method 9. However, Golgi staining is considered to be unpredictable. Indeed, groups of nerve cells and fiber tracts are labeled randomly, with particular areas often appearing completely stained while adjacent areas are devoid of staining. Recent studies have shown that IUE of fluorescent constructs represents an attractive alternative method to study dendrites, spines as well as synapses in mutant / wild-type mice 10-11 (Figure 1A). Moreover in comparison to the generation of mouse knockouts, IUE represents a rapid approach to perform gain and loss of function studies in specific population of cells during a specific time window. In addition, IUE has been successfully used with inducible gene expression or inducible RNAi approaches to refine the temporal control over the expression of a gene or shRNA 12. These advantages of IUE have thus opened new dimensions to study the effect of gene expression/suppression on dendrites and spines not only in specific cerebral structures (Figure 1B) but also at a specific time point of development (Figure 1C).
Finally, IUE provides a useful tool to identify functional interactions between genes involved in dendrite, spine and/or synapse development. Indeed, in contrast to other gene transfer methods such as virus, it is straightforward to combine multiple RNAi or transgenes in the same population of cells. 
In summary, IUE is a powerful method that has already contributed to the characterization of molecular mechanisms underlying brain function and disease and it should also be useful in the study of dendrites and spines.

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

This article describes in detail a protocol to electroporate in utero the cerebral cortex and the hippocampus at E14.5 in mice. We also show that this is a valuable method to study dendrites and spines in these two cerebral regions.

Visualização e manipulação genética de dendritos e espinhas no córtex cerebral e hipocampo do rato usando<em&gt; In utero</em&gt; Eletroporação

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date ...

Stereotaxic Injection of a Viral Vector for Conditional Gene Manipulation in the Mouse Spinal Cord

Intraparenchymal injection of a viral vector enables conditional gene manipulation in distinct populations of neurons or particular regions of the central nervous system. We demonstrate a stereotaxic injection technique that allows targeted gene expression or silencing in the dorsal horn of the mouse spinal cord. The surgical procedure is brief. It requires laminectomy of a single vertebra, providing for quick recovery of the animal and unimpaired motility of the spine. Controlled injection of a small vector suspension volume at low speed and use of a microsyringe with beveled glass cannula minimize the tissue lesion. The local immune response to the vector depends on the intrinsic properties of the virus employed; in our experience, it is minor and short-lived when a recombinant adeno-associated virus is used. A reporter gene such as enhanced green fluorescent protein facilitates monitoring spatial distribution of the vector, and the efficacy and cellular specificity of the transfection.

Viral vectors allow for targeted gene manipulation. We demonstrate a method for conditional gene expression or ablation in the mouse spinal cord, using stereotaxic injection of a viral vector into the dorsal horn, a prominent site of synaptic contact between primary somatosensory afferents and neurons of the central nervous system.

Injeção estereotáxica de um vetor viral para manipulação genética condicional no Cabo de Rato Spinal

Intraparenchymal injection of a viral vector enables conditional gene manipulation in distinct populations of neurons or particular regions of the central ...

Building An Open-source Robotic Stereotaxic Instrument

This protocol includes the designs and software necessary to upgrade an existing stereotaxic instrument to a robotic (CNC) stereotaxic instrument for around $1,000 (excluding a drill), using industry standard stepper motors and CNC controlling software. Each axis has variable speed control and may be operated simultaneously or independently. The robot&#39;s flexibility and open coding system (g-code) make it capable of performing custom tasks that are not supported by commercial systems. Its applications include, but are not limited to, drilling holes, sharp edge craniotomies, skull thinning, and lowering electrodes or cannula. In order to expedite the writing of g-coding for simple surgeries, we have developed custom scripts that allow individuals to design a surgery with no knowledge of programming. However, for users to get the most out of the motorized stereotax, it would be beneficial to be knowledgeable in mathematical programming and G-Coding (simple programming for CNC machining).
The recommended drill speed is greater than 40,000 rpm. The stepper motor resolution is 1.8&#176;/Step, geared to 0.346&#176;/Step. A standard stereotax has a resolution of 2.88 &#956;m/step. The maximum recommended cutting speed is 500 &#956;m/sec. The maximum recommended jogging speed is 3,500 &#956;m/sec. The maximum recommended drill bit size is HP 2.

This protocol includes the designs and software necessary to upgrade an existing stereotaxic instrument to a robotic (computer numeric controlled; CNC) stereotaxic instrument for around $1,000 (excluding a drill).

Construir um Robotic estereotáxica Instrumento Open-source

This protocol includes the designs and software necessary to upgrade an existing stereotaxic instrument to a robotic (CNC) stereotaxic instrument for ...

Mouse Microsurgery Infusion Technique for Targeted Substance Delivery into the CNS via the Internal Carotid Artery

Animal models of central nervous system (CNS) diseases and, consequently, blood-brain barrier disruption diseases, require the delivery of exogenous substances into the brain. These exogenous substances may induce injurious impact or constitute therapeutic strategy. The most common delivery methods of exogenous substances into the brain are based on systemic deliveries, such as subcutaneous or intravenous routes. Although commonly used, these approaches have several limitations, including low delivery efficacy into the brain. In contrast, surgical methods that locally deliver substances into the CNS are more specific and prevent the uptake of the exogenous substances by other organs. Several surgical methods for CNS delivery are available; however, they tend to be very traumatic. Here, we describe a mouse infusion microsurgery technique, which effectively delivers substances into the brain via the internal carotid artery, with minimal trauma and no interference with normal CNS functionality.

Mouse Microsurgery Infusion Technique for Targeted Substance Delivery into the CNS via the Internal Carotid Artery

The present protocol describes a mouse microsurgery infusion technique, which effectively delivers substances directly into the brain via the internal carotid artery.

Técnica do mouse Microcirurgia de infusão para distribuição de substâncias alvejado no SNC<i&gt; via</i&gt; Artéria carótida interna

Animal models of central nervous system (CNS) diseases and, consequently, blood-brain barrier disruption diseases, require the delivery of exogenous ...

Stereotaxic Surgery for Genetic Manipulation in Striatal Cells of Neonatal Mouse Brains

Many genes are expressed in embryonic brains, and some of them are continuously expressed in the brain after birth. For such persistently expressed genes, they may function to regulate the developmental process and/or physiological function in neonatal brains. To investigate neurobiological functions of specific genes in the brain, it is essential to inactivate genes in the brain. Here, we describe a simple stereotaxic method to inactivate gene expression in the striatum of transgenic mice at neonatal time windows. AAV-eGFP-Cre viruses were microinjected into the striatum of Ai14 reporter gene mice at postnatal day (P) 2 by stereotaxic brain surgery. The tdTomato reporter gene expression was detected in P14 striatum, suggesting a successful Cre-loxP mediated DNA recombination in AAV-transduced striatal cells. We further validated this technique by microinjecting AAV-eGFP-Cre viruses into P2Foxp2fl/fl mice. Double labeling of GFP and Foxp2 showed that GFP-positive cells lacked Foxp2 immunoreactivity in P9 striatum, suggesting the loss of Foxp2 protein in AAV-eGFP-Cre transduced striatal cells. Taken together, these results demonstrate an effective genetic deletion by stereotaxically microinjected AAV-eGFP-Cre viruses in specific neuronal populations in the neonatal brains of floxed transgenic mice. In conclusion, our stereotaxic technique provides an easy and simple platform for genetic manipulation in neonatal mouse brains. The technique can not only be used to delete genes in specific regions of neonatal brains, but it also can be used to inject pharmacological drugs, neuronal tracers, genetically modified optogenetics and chemogenetics proteins, neuronal activity indicators and other reagents into the striatum of neonatal mouse brains.

We describe a protocol of stereotaxic surgery with a homemade head-fixed device for microinjecting reagents into the striatum of neonatal mouse brains. This technique allows genetic manipulation in neuronal cells of specific regions of neonatal mouse brains.

Cirurgia estereotáxica para manipulação genética em células estriadas de cérebro de rato Neonatal

Many genes are expressed in embryonic brains, and some of them are continuously expressed in the brain after birth. For such persistently expressed genes, ...

Recording Spatially Restricted Oscillations in the Hippocampus of Behaving Mice

The local field potential (LFP) emerges from ion movements across neural membranes. Since the voltage recorded by LFP electrodes reflects the summed electrical field of a large volume of brain tissue, extracting information about local activity is challenging. Studying neuronal microcircuits, however, requires a reliable distinction between truly local events and volume-conducted signals originating in distant brain areas. Current source density (CSD) analysis offers a solution for this problem by providing information about current sinks and sources in the vicinity of the electrodes. In brain areas with laminar cytoarchitecture such as the hippocampus, one-dimensional CSD can be obtained by estimating the second spatial derivative of the LFP. Here, we describe a method to record multilaminar LFPs using linear silicon probes implanted into the dorsal hippocampus. CSD traces are computed along individual shanks of the probe. This protocol thus describes a procedure to resolve spatially restricted neuronal network oscillations in the hippocampus of freely moving mice.

This protocol describes the recording of local field potentials with multi-shank linear silicon probes. Conversion of the signals using current source density analysis allows the reconstruction of local electrical activity in the mouse hippocampus. With this technique, spatially restricted brain oscillations can be studied in freely moving mice.

Gravação espacialmente restrita oscilações no hipocampo de ratos de comportar-se

The local field potential (LFP) emerges from ion movements across neural membranes. Since the voltage recorded by LFP electrodes reflects the summed ...

Transplantation of Chemogenetically Engineered Cortical Interneuron Progenitors into Early Postnatal Mouse Brains

Neuronal development is regulated by a complex combination of environmental and genetic factors. Assessing the relative contribution of each component is a complicated task, which is particularly difficult in regards to the development of &#947;-aminobutyric acid (GABA)ergic cortical interneurons (CIs). CIs are the main inhibitory neurons in the cerebral cortex, and they play key roles in neuronal networks, by regulating both the activity of individual pyramidal neurons, as well as the oscillatory behavior of neuronal ensembles. They are generated in transient embryonic structures (medial and caudal ganglionic eminences -&#160;MGE and CGE) that are very difficult to efficiently target using in utero electroporation approaches. Interneuron progenitors migrate long distances during normal embryonic development, before they integrate in the cortical circuit. This remarkable ability to disperse and integrate into a developing network can be hijacked by transplanting embryonic interneuron precursors into early post-natal host cortices. Here, we present a protocol that allows genetic modification of embryonic interneuron progenitors using focal ex vivo electroporation. These engineered interneuron precursors are then transplanted into early post-natal host cortices, where they will mature into easily identifiable CIs. This protocol allows the use of multiple genetically encoded tools, or the ability to regulate the expression of specific genes in interneuron progenitors, in order to investigate the impact of either genetic or environmental variables on the maturation and integration of CIs.

Here we present a protocol, designed to use chemogenetic tools to manipulate the activity of cortical interneuron progenitors transplanted into the cortex of early postnatal mice.

Transplantação dos progenitors corticais Quimiogenetically projetados do interneuron em cérebros Postnatal adiantados do rato

Neuronal development is regulated by a complex combination of environmental and genetic factors. Assessing the relative contribution of each component is ...

Stereotaxic Surgery for Implantation of Microelectrode Arrays in the Common Marmoset (Callithrix jacchus)

Marmosets (Callithrix jacchus) are small non-human primates that are gaining popularity in biomedical and preclinical research, including the neurosciences. Phylogenetically, these animals are much closer to humans than rodents. They also display complex behaviors, including a wide range of vocalizations and social interactions. Here, an effective stereotaxic neurosurgical procedure for implantation of recording electrode arrays in the common marmoset is described. This protocol also details the pre- and postoperative steps of animal care that are required to successfully perform such a surgery. Finally, this protocol shows an example of local field potential and spike activity recordings in a freely behaving marmoset 1 week after the surgical procedure. Overall, this method provides an opportunity to study the brain function in awake and freely behaving marmosets. The same protocol can be readily used by researchers working with other small primates. In addition, it can be easily modified to allow other studies requiring implants, such as stimulating electrodes, microinjections, implantation of optrodes or guide cannulas, or ablation of discrete tissue regions.

Stereotaxic Surgery for Implantation of Microelectrode Arrays in the Common Marmoset Callithrix jacchus

This work presents a protocol to perform a stereotaxic, neurosurgical implantation of&#160;microelectrode arrays in the common marmoset. This method specifically enables electrophysiological recordings in freely behaving animals but can be easily adapted to any other similar neurosurgical intervention in this species (e.g., cannula for drug administration or electrodes for brain stimulation).