The authors acknowledge the help from the staff of Laboratory Animal Medicine, University of Nevada, Reno. MM received funding from Nevada INBRE.

El protocolo (Número-00682) se describe a continuación es aprobado por el Comité Institucional de Cuidado y Uso de Animales (IACUC) de la Universidad de Nevada, Reno y sigue las directrices de la Universidad de Nevada, animal comité de ética de investigación de Reno. Brevemente, los ratones fueron anestesiados con isoflurano y se utilizó un cono de la nariz para mantener los niveles de isoflurano continuas durante 20 min. Un ungüento veterinario se utilizó para evitar el secado de los ojos bajo anestesia. pellizco del dedo del pie se utilizó para determinar el nivel de anestesia y la frecuencia respiratoria se midió durante todo el procedimiento. Los ratones se mantuvieron en jaulas individuales y supervisados ​​hasta que se recuperó la conciencia. Los ratones se usaron sólo una vez para marcar la alimentación y se sacrificaron recuperación posterior garrapata. La eutanasia se llevó a cabo por el Laboratorio de personal del personal de medicamentos para animales. CO 2 y dislocación cervical se utilizaron para la eutanasia de los animales. NOTA: Trabajar sobre las garrapatas requiere el uso de la ecuación de protección personal completaUipment. Para contar las garrapatas inmaduras, use batas de laboratorio blancas, guantes largos de nitrilo para cubrir la abertura de la manga de la bata de laboratorio tapada con gomas y zapatos cerrados. Para infestar ratones con garrapatas, utilice una red de pelo, trajes desechables, guantes de manga larga sobre las mangas y cubiertas de los pies. Use batas blancas o de colores claros para detectar garrapatas rebeldes. Revise periódicamente los guantes y las mangas de las garrapatas. 1. Sala de Preparación para la Vivienda de Animales <ol><li> Designe una sala separada para alojar ratones infestados con garrapatas. Coloque una alfombra pegajosa o cinta adhesiva de dos caras en el exterior y en el interior de la puerta para evitar el escape accidental de garrapatas. </li></ol> 2. Cuenta de las garrapatas para la infestación de ratón <ol><li> Coloque cinta adhesiva de doble cara alrededor de los bordes superiores de un recipiente de plástico de 7 &quot;x 5&quot; x 14 &quot;y rellene con agua hasta aproximadamente 2 cm. </li><li> Coloque otra pequeña caja o placa de Petri en el centro del recipiente y rellene con agua hasta 1 cm,# 34; foso&quot; alrededor de este envase </li><li> Almacenar el vial que contiene larvas o ninfas dentro de la placa de Petri. </li><li> Utilice un pincel fino para eliminar las larvas o ninfas del vial y contar con el microscopio. Contar 50 larvas o ninfas 25 en viales de centelleo separados. Cubrir los viales inmediatamente con malla de nylon o pantalla de tela organdí y cerrar con bandas de goma. </li></ol> 3. Los ratones que infestan con garrapatas inmaduras <ol><li> Utilizar un banco de trabajo de color blanco o claro y se adhieren cinta de doble cara en todo el perímetro de la zona de trabajo. </li><li> Anestesiar al ratón con isoflurano. Comprobar el nivel de anestesia mediante la pizca dedo del pie. Una vez anestesiados, transferir el ratón para una resistencia de calentamiento cubierto con toallas de papel y adjuntar a un cono de la nariz para el suministro continuo de isoflurano. </li><li> Aplicar petróleo ungüento para los ojos a base de jalea para evitar la sequedad. Tenga en cuenta el patrón de respiración del ratón para ajustar los niveles de isoflurano (80 -. 230 respiraciones por min es normal, Reducir el nivel de isoflurano si brtasa eathing es de menos de 80 respiraciones por min para evitar la muerte del animal). </li><li> Tomar un vial de 50 larvas o ninfas 25 y el lugar garrapatas bajo la piel en la cabeza entre las orejas con un cepillo de pintura (Figura 1). Mantenga ratón bajo anestesia durante 20 min después de la colocación para dar tiempo suficiente para que las garrapatas adjuntar. </li><li> Mueve el ratón para un plástico caja de zapatos en forma de jaula de ratón estándar desechable con tapas estáticas y ropa de cama blanco. Proporcionar juguetes, agua y alimento ad libitum de acuerdo con el cuidado normal de ratón. </li><li> Almacenar la jaula del ratón dentro de una jaula de rata o jerbo grande forrado con cinta de doble cara adhesiva alrededor de los bordes superiores. Llene la jaula externa con 3 cm de agua (Figura 2). </li></ol> 4. Recogida de las garrapatas de ratones <ol><li> garrapatas inmaduras se separan entre los días 3 y 6 de la alimentación. Compruebe las jaulas y un foso de agua de garrapatas saciadas todos los días después del día 3. </li><li> Recoger las garrapatas desprendidas de la jaula entre los días 4 y 6. Uso de un dolorT pincel o fórceps suave para recoger las garrapatas congestionadas y almacenar en viales de centelleo limpios cubiertos con tela de malla de nylon asegurada con bandas de goma. </li><li> Anestesiar y comprobar el ratón en el día 7 para cualquier resto de garrapatas adjunta. </li><li> Compruebe la ropa de cama, el comedero y la botella de agua para las garrapatas congestionadas el día 7. Eutanee los ratones siguiendo el procedimiento descrito anteriormente. Autoclave las jaulas desechables, la ropa de cama, la botella de agua, y el comedero de la comida para evitar cualquier garrapatas sin escape. </li></ol> 5. Almacenamiento de garrapatas Fed <ol><li> Mantener las garrapatas con congestión a 90% de humedad, 20 ° C y un ciclo de 12:12 luz: oscuridad en una incubadora de humedad y temperatura controlada hasta que ocurre la muda. Esto puede tomar aproximadamente 12 a 18 semanas para ocurrir. </li><li> Mientras que la alta humedad promueve la supervivencia de la garrapata, también hace que las garrapatas sean propensas al crecimiento excesivo de hongos. Compruebe las garrapatas alimentadas bajo el microscopio al menos una vez a la semana para el moho. Si se detecta, lave a los supervivientes en etanol al 70% durante 5min., enjuague en agua, la transferencia a un papel de filtro para secar, y transferir a las nuevas, viales limpios. </li></ol>

<ol>
	<li>Gleim, E. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Factors+associated+with+tick+bites+and+pathogen+prevalence+in+ticks+parasitizing+humans+in+Georgia,+USA.">Factors associated with tick bites and pathogen prevalence in ticks parasitizing humans in Georgia, USA.</a> Parasites &amp;amp; Vectors. 9 (125), 1-13 (2016).</li><li>Krober, T., Guerin, P. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+feeding+assays+for+hard+ticks.">In vitro feeding assays for hard ticks.</a> Trends Parasitol. 23 (9), 445-449 (2007).</li><li>Kuhnert, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Feeding+of+Hard+Ticks+In+Vitro:+New+Perspectives+for+Rearing+and+for+the+Identification+of+Systemic+Acaricides.">Feeding of Hard Ticks In Vitro: New Perspectives for Rearing and for the Identification of Systemic Acaricides.</a> ALTEX. 13 (2), 76-87 (1996).</li><li>Voigt, W. P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+feeding+of+instars+of+the+ixodid+tick+Amblyomma+variegaturn+on+skin+membranes+and+its+application+to+the+transmission+of+Theileria+mutans+and+Cowdria+ruminantium.">In vitro feeding of instars of the ixodid tick Amblyomma variegaturn on skin membranes and its application to the transmission of Theileria mutans and Cowdria ruminantium.</a> Parasitol. 107, 257-263 (1993).</li><li>Gregson, J. D., Smith, C. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ticks.">Ticks.</a> Insect Colonization and Mass Production. , 49-72 (1966).</li><li>Sonenshine, D. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Biology of Ticks. 2, (1993).</li><li>Bouchard, K. R., Wikel, S. K., Marquaedt, W. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Care,+maintenance,+and+experimental+infestation+of+ticks+in+the+laboratory+setting.">Care, maintenance, and experimental infestation of ticks in the laboratory setting.</a> Biology of Disease Vectors. , 705-711 (2005).</li><li>Schumaker, T. S., Barros, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Life+cycle+of+Ornithodoros+(Alectorobius)+talaje.+(Acari:Argasidae)+in+laboratory.">Life cycle of Ornithodoros (Alectorobius) talaje. (Acari:Argasidae) in laboratory.</a> J Med Entomol. 32, 249-254 (1995).</li><li>Banks, C. W., Oliver, J. H., Hopla, C. E., Dotson, E. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Laboratory+life+cycle+of+Ixodes+woodi+(Acari:Ixodidae).">Laboratory life cycle of Ixodes woodi (Acari:Ixodidae).</a> J. Med. Entomol. 35, 177-179 (1998).</li><li>James, A. M., Oliver, J. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Feeding+and+host+preference+of+immature+Ixodes+dammini,I.scapularis,and+I.pacificus.(Acari:Ixodidae).">Feeding and host preference of immature Ixodes dammini,I.scapularis,and I.pacificus.(Acari:Ixodidae).</a> J. Med. Entomol. 27, 324-330 (1990).</li></ol>

The authors have nothing to disclose.

Los pasos críticos dentro del Protocolo Es importante tener varios niveles de medidas de seguridad cuando la crianza de garrapatas para evitar el escape accidental. El uso de cinta adhesiva y un foso de agua son fundamentales para garantizar la seguridad. Es importante mantener el ratón anestesiado en una almohadilla de calefacción para mantener constante la temperatura del cuerpo. También se encontró que afeitarse el ratón no proporciona ningún beneficio adicional para la fijación de l...

Las garrapatas son ectoparásitos hematófagos obligados de los vertebrados y se distribuyen en todo el mundo. En los Estados Unidos, al menos 11 especies de garrapatas son vectores de patógenos de importancia para la salud pública 1. Ixodes scapularis es responsable de la transmisión de varios patógenos tales como los agentes causantes de la enfermedad de Lyme (Borrelia burgdorferi) fiebre recurrente (B. miyamotoi), anaplasmosis granulocítica humana (Anaplasma phagocytophilum), y babesiosis (Babesia spp.). A pesar de la importancia de I. scapularis como vector de la enfermedad, la recogida de estos arácnidos en abundancia en la naturaleza para los estudios en el laboratorio no siempre es factible. Por lo tanto, la producción de garrapatas sanos criados en el laboratorio es esencial para los estudios sobre la biología de la señal, y de la señal de patógenos interacciones. El ciclo vital de todas las garrapatas duras (Ixodidae) de la familia, incluyendo I. scapularis, consiste en el huevo y tres activetapas e: larva, ninfa y adulto. Cada etapa activa se alimenta de un huésped vertebrado. Las complejas interacciones que tienen lugar entre las garrapatas y sus anfitriones lo largo de varios días de fijación y la alimentación son casi imposibles de replicar usando alimentadores artificiales, y es improbable que proporcionar suficientes números de Fed garrapatas para la cría masiva de 2, 3, 4. Por lo tanto, los ratones y conejos vivos son los más usados ​​como anfitriones para la crianza de inmaduros (larvas y ninfas), y etapas maduras (adultos) de las garrapatas, respectivamente. El requisito de múltiples hosts para la alimentación de sangre durante cada etapa de desarrollo complica la cría de garrapata, y es el tiempo y el coste intensivo de 5, 6, 7. La mayoría de los protocolos de garrapatas cría requieren mantenimiento de los ratones en una jaula de alambre suelo de rejilla suspendido 7, 8 o en una ca cilíndricaGe de tales dimensiones que el animal no puede moverse libremente y prepararse 6 , 9 , 10 . Estas jaulas cilíndricas se transfieren posteriormente a una jaula de caja de zapatos con una rejilla de alambre. Las garrapatas engorged, separadas se recogen entonces del agua debajo. Sin embargo, este método resulta en la exposición de las garrapatas alimentadas a agua contaminada con orina y heces que pueden aumentar el crecimiento de hongos y la mortalidad de garrapatas 9 . Además, aumenta la posibilidad de escape de garrapata de la depresión de agua, así como causar estrés a los ratones. Para eludir estos problemas, aquí se demuestra la alimentación de garrapatas de larvas en ratones dentro de jaulas desechables de tipo caja de zapatos de plástico. Este método permite el comportamiento normal de los ratones, aumenta la recuperación de garrapatas engorged, y disminuye la mortalidad de garrapatas debido a la contaminación.

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rearing ixodes scapularis, the black-legged tick: feeding immature stages on mice

Ixodes scapularis , el vector de la enfermedad de Lyme, es uno de los más importantes vectores de enfermedades en el este y el medio oeste de los Estados Unidos. Esta especie es una garrapata de tres huéspedes que requiere una comida de sangre de un huésped vertebrado para cada etapa de desarrollo, y las hembras adultas requieren una comida de sangre para su reproducción. Las garrapatas larvales se unen a su huésped durante 3 - 5 días para la alimentación y dejar al huésped cuando están completamente hinchadas. Esta dependencia de varios huéspedes diferentes y el largo tiempo de fijación para el engorgamiento complica la elevación de las garrapatas en el laboratorio. Sin embargo, para entender la biología de la garrapata y las interacciones de la garrapata-patógeno, la producción de garrapatas sanas, criadas en laboratorio es esencial. Aquí, demostramos un sencillo, costo-efectiva protocolo para inmaduros tick alimentación en ratones. Se modificaron los protocolos existentes para disminuir el estrés en los ratones y el aumento del éxito de la alimentación de garrapatas y la supervivencia mediante el uso de jaulas desechables sin fondo de malla para evitar el contacto de garrapatas wiAgua contaminada con orina y heces de ratones.

Hemos modificado los protocolos de cría de garrapatas existentes 6, 10 para mejorar la eficiencia de la alimentación y la reducción del estrés en el host del ratón. Los resultados demuestran que las jaulas de ratón estándar estilo caja de zapatos son muy adecuados para la cría de garrapata. La ropa de cama blanca proporciona un buen contraste para facilitar la recogida de las garrapatas alimentadas. La mayoría de las gar...

Watch this Scientific Journal Video about Rearing Ixodes scapularis, the Black-legged Tick: Feeding Immature Stages on Mice at JoVE.com

Rearing Ixodes scapularis, the Black-legged Tick: Feeding Immature Stages on Mice

La producción de garrapatas sanos criados en el laboratorio es esencial para los estudios sobre la biología de la señal, y de la señal de patógenos interacciones. Aquí se demuestra un protocolo sencillo para la alimentación de la garrapata inmadura que es rentable y menos estresante para ratones.

La cría<em&gt; Ixodes scapularis,</em&gt; La garrapata de patas Negro: La alimentación de los estados inmaduros de ratones

Ixodes scapularis, the vector of Lyme disease, is one of the most important disease vectors in the eastern and Midwestern United States. This species is a ...

rearing-ixodes-scapularis-black-legged-tick-feeding-immature-stages

Research

JoVE Journal

Immunology and Infection

Rearing Ixodes scapularis, the Black-legged Tick: Feeding Immature Stages on Mice

11.9K vistas.  University of Nevada, Reno.  La producción de garrapatas sanos criados en el laboratorio es esencial para los estudios sobre la biología de la señal, y de la señal de patógenos interacciones. Aquí se demuestra un protocolo sencillo para la alimentación de la garrapata inmadura que es rentable y menos estresante para ratones. 

Video: Rearing Ixodes scapularis, the Black-legged Tick: Feeding Immature Stages on Mice

Methods for Rapid Transfer and Localization of Lyme Disease Pathogens Within the Tick Gut

Lyme disease is caused by infection with the spirochete pathogen Borrelia burgdorferi, which is maintained in nature by a tick-rodent infection cycle 1. A tick-borne murine model 2 has been developed to study Lyme disease in the laboratory. While na&iacute;ve ticks can be infected with B. burgdorferi by feeding them on infected mice, the molting process takes several weeks to months to complete. Therefore, development of more rapid and efficient tick infection techniques, such as a microinjection-based procedure, is an important tool for the study of Lyme disease 3,4. The procedure requires only hours to generate infected ticks and allows control over the delivery of equal quantities of spirochetes in a cohort of ticks. This is particularly important as the generation of B. burgdorferi infected ticks by the natural feeding process using mice fails to ensure 100% infection rate and potentially results in variation of pathogen burden amongst fed ticks. Furthermore, microinjection can be used to infect ticks with B. burgdorferi isolates in cases where an attenuated strain is unable to establish infection in mice and thus can not be naturally acquired by ticks 5. This technique can also be used to deliver a variety of other biological materials into ticks, for example, specific antibodies or double stranded RNA 6. In this article, we will demonstrate the microinjection of nymphal ticks with in vitro-grown B. burgdorferi. We will also describe a method for localization of Lyme disease pathogens in the tick gut using confocal immunofluorescence microscopy.

Lyme disease research studies often require generation of ticks infected with the pathogen Borrelia burgdorferi, a process that typically takes several weeks. Here we demonstrate a microinjection-based tick infection procedure that can be accomplished within hours. We also demonstrate an immunofluorescence method for in situ localization of B. burgdorferi within ticks.

Métodos para la transferencia rápida y localización de la enfermedad de Lyme patógenos en el intestino garrapatas

Lyme disease is caused by infection with the spirochete pathogen Borrelia burgdorferi, which is maintained in nature by a tick-rodent infection cycle 1.  ...

Investigación

Inmunología e Infección

Saliva, Salivary Gland, and Hemolymph Collection from Ixodes scapularis Ticks

Ticks are found worldwide and afflict humans with many tick-borne illnesses. Ticks are vectors for pathogens that cause Lyme disease and tick-borne relapsing fever (Borrelia spp.), Rocky Mountain Spotted fever (Rickettsia rickettsii), ehrlichiosis (Ehrlichia chaffeensis and E. equi), anaplasmosis (Anaplasma phagocytophilum), encephalitis (tick-borne encephalitis virus), babesiosis (Babesia spp.), Colorado tick fever (Coltivirus), and tularemia (Francisella tularensis) 1-8. To be properly transmitted into the host these infectious agents differentially regulate gene expression, interact with tick proteins, and migrate through the tick 3,9-13. For example, the Lyme disease agent, Borrelia burgdorferi, adapts through differential gene expression to the feast and famine stages of the tick's enzootic cycle 14,15. Furthermore, as an Ixodes tick consumes a bloodmeal Borrelia replicate and migrate from the midgut into the hemocoel, where they travel to the salivary glands and are transmitted into the host with the expelled saliva 9,16-19.
As a tick feeds the host typically responds with a strong hemostatic and innate immune response 11,13,20-22. Despite these host responses, I. scapularis can feed for several days because tick saliva contains proteins that are immunomodulatory, lytic agents, anticoagulants, and fibrinolysins to aid the tick feeding 3,11,20,21,23. The immunomodulatory activities possessed by tick saliva or salivary gland extract (SGE) facilitate transmission, proliferation, and dissemination of numerous tick-borne pathogens 3,20,24-27. To further understand how tick-borne infectious agents cause disease it is essential to dissect actively feeding ticks and collect tick saliva. This video protocol demonstrates dissection techniques for the collection of hemolymph and the removal of salivary glands from actively feeding I. scapularis nymphs after 48 and 72 hours post mouse placement. We also demonstrate saliva collection from an adult female I. scapularis tick.

Saliva, Salivary Gland, and Hemolymph Collection from Ixodes scapularis Ticks

The collection of infected tick hemolymph, salivary glands, and saliva is important to study how tick-borne pathogens cause disease. In this protocol we demonstrate how to collect hemolymph and salivary glands from feeding Ixodes scapularis nymphs. We also demonstrate saliva collection from female I. scapularis adults.

La saliva, la glándula salival, y la Colección hemolinfa de las garrapatas Ixodes scapularis

Ticks are found worldwide and afflict humans with many tick-borne illnesses. Ticks are vectors for pathogens that cause Lyme disease and tick-borne ...

Rearing and Injection of Manduca sexta Larvae to Assess Bacterial Virulence

Manduca sexta, commonly known as the tobacco hornworm, is considered a significant agricultural pest, feeding on solanaceous plants including tobacco and tomato. The susceptibility of M. sexta larvae to a variety of entomopathogenic bacterial species1-5, as well as the wealth of information available regarding the insect's immune system6-8, and the pending genome sequence9 make it a good model organism for use in studying host-microbe interactions during pathogenesis. In addition, M. sexta larvae are relatively large and easy to manipulate and maintain in the laboratory relative to other susceptible insect species. Their large size also facilitates efficient tissue/hemolymph extraction for analysis of the host response to infection.
The method presented here describes the direct injection of bacteria into the hemocoel (blood cavity) of M. sexta larvae. This approach can be used to analyze and compare the virulence characteristics of various bacterial species, strains, or mutants by simply monitoring the time to insect death after injection. This method was developed to study the pathogenicity of Xenorhabdus and Photorhabdus species, which typically associate with nematode vectors as a means to gain entry into the insect. Entomopathogenic nematodes typically infect larvae via natural digestive or respiratory openings, and release their symbiotic bacterial contents into the insect hemolymph (blood) shortly thereafter10. The injection method described here bypasses the need for a nematode vector, thus uncoupling the effects of bacteria and nematode on the insect. This method allows for accurate enumeration of infectious material (cells or protein) within the inoculum, which is not possible using other existing methods for analyzing entomopathogenesis, including nicking11 and oral toxicity assays12. Also, oral toxicity assays address the virulence of secreted toxins introduced into the digestive system of larvae, whereas the direct injection method addresses the virulence of whole-cell inocula.
The utility of the direct injection method as described here is to analyze bacterial pathogenesis by monitoring insect mortality. However, this method can easily be expanded for use in studying the effects of infection on the M. sexta immune system. The insect responds to infection via both humoral and cellular responses. The humoral response includes recognition of bacterial-associated patterns and subsequent production of various antimicrobial peptides7; the expression of genes encoding these peptides can be monitored subsequent to direct infection via RNA extraction and quantitative PCR13. The cellular response to infection involves nodulation, encapsulation, and phagocytosis of infectious agents by hemocytes6. To analyze these responses, injected insects can be dissected and visualized by microscopy13, 14.

Rearing and Injection of Manduca sexta Larvae to Assess Bacterial Virulence

The method described here utilizes direct injection of entomopathogenic bacteria into the hemocoel of Manduca sexta insect larvae. M. sexta is a commercially available and well-studied insect. Thus, this method represents a simple approach to analyzing host-bacterial interactions from the perspective of one or both partners.

Crianza y la inyección de<em&gt; Manduca sexta</em&gt; Las larvas para evaluar la virulencia bacteriana

Manduca sexta, commonly known as the tobacco hornworm, is considered a significant agricultural pest, feeding on solanaceous plants including tobacco and ...

Feeding of Ticks on Animals for Transmission and Xenodiagnosis in Lyme Disease Research

Transmission of the etiologic agent of Lyme disease, Borrelia burgdorferi, occurs by the attachment and blood feeding of Ixodes species ticks on mammalian hosts. In nature, this zoonotic bacterial pathogen may use a variety of reservoir hosts, but the white-footed mouse (Peromyscus leucopus) is the primary reservoir for larval and nymphal ticks in North America. Humans are incidental hosts most frequently infected with B. burgdorferi by the bite of ticks in the nymphal stage. B. burgdorferi adapts to its hosts throughout the enzootic cycle, so the ability to explore the functions of these spirochetes and their effects on mammalian hosts requires the use of tick feeding. In addition, the technique of xenodiagnosis (using the natural vector for detection and recovery of an infectious agent) has been useful in studies of cryptic infection. In order to obtain nymphal ticks that harbor B. burgdorferi, ticks are fed live spirochetes in culture through capillary tubes. Two animal models, mice and nonhuman primates, are most commonly used for Lyme disease studies involving tick feeding. We demonstrate the methods by which these ticks can be fed upon, and recovered from animals for either infection or xenodiagnosis.

Lyme disease is the most commonly-reported vector-borne disease in North America. The causative agent, Borrelia burgdorferi is a spirochete bacterium transmitted by Ixodid ticks. Transmission and detection of infection in animal models is optimized by the use of tick feeding, which we describe here.

La alimentación de las garrapatas de los animales para Transmisión y xenodiagnóstico en Lyme Investigación de Enfermedades

Transmission of the etiologic agent of Lyme disease, Borrelia burgdorferi, occurs by the  attachment and blood feeding of Ixodes species ticks on ...

Measuring Local Anaphylaxis in Mice

Allergic responses are the result of the activation of mast cells and basophils, and the subsequent release of vasoactive and proinflammatory mediators. Exposure to an allergen in a sensitized individual can result in clinical symptoms that vary from minor erythema to life threatening anaphylaxis. In the laboratory, various animal models have been developed to understand the mechanisms driving allergic responses. Herein, we describe a detailed method for measuring changes in vascular permeability to quantify localized allergic responses. The local anaphylaxis assay was first reported in the 1920s, and has been adapted from the technique published by Kojima et al. in 20071. In this assay, mice sensitized to OVA are challenged in the left ear with vehicle and in the right ear with OVA. This is followed by an intravenous injection of Evans Blue dye. Ten min after injecting Evans Blue, the animal is euthanized and the dye that has extravasated into the ears is extracted overnight in formamide. The absorbance of the extracted dye is then quantified with a spectrophotometer. This method reliably results in a visual and quantifiable manifestation of a local allergic response.

Allergic responses, characterized by the activation of mast cells and basophils, are driven by the cross-linking of IgE and release of proinflammatory mediators. A quantitative assessment of allergic responses can be achieved by using Evans Blue dye to monitor changes in vascular permeability after allergen challenge.

Medición anafilaxia local en ratones

Allergic responses are the result of the activation of mast cells and basophils, and the subsequent release of vasoactive and proinflammatory mediators. ...

Generation of Immature, Mature and Tolerogenic Dendritic Cells with Differing Metabolic Phenotypes

Immune response results from a complex interplay between the antigen non-specific innate immune system and the antigen specific adaptive immune system. The immune system is a constant balance in maintaining tolerance to self-molecules and reacting rapidly to pathogens. Dendritic cells (DCs) are powerful professional antigen presenting cells that link the innate immune system to the adaptive immune system and balance the adaptive response between self and non-self. Depending on the maturation signals, immature dendritic cells can be selectively stimulated to differentiate into immunogenic or tolerogenic DCs. Immunogenic dendritic cells provide proliferation signals to antigen-specific T cells for clonal expansion; while tolerogenic dendritic cells regulate tolerance by antigen-specific T-cell deletion or clonal expansion of regulatory T-cells. Due to this unique property, dendritic cells are highly sought after as therapeutic agents for cancer and autoimmune diseases. Dendritic cells can be loaded with specific antigens in vitro and injected into the human body to mount a specific immune response both immunogenic and tolerogenic. This work presents a means to generate in vitro from monocytes, immature monocyte derived dendritic cells (moDCs), tolerogenic and mature moDCs that differ in surface marker expression, function and metabolic phenotypes.

Immature dendritic cells can be selectively differentiated into tolerogenic or mature dendritic cells to regulate the balance between immunity and tolerance. This work presents a means to generate from immature monocyte derived dendritic cells (moDCs), in vitro tolerogenic and mature moDCs that differ in metabolic phenotypes.

Generación de inmaduras, maduras y tolerogénico células dendríticas con diferentes fenotipos metabólico

Immune response results from a complex interplay between the antigen non-specific innate immune system and the antigen specific adaptive immune system. ...

Development of a Hepatitis B Virus Reporter System to Monitor the Early Stages of the Replication Cycle

Currently, it is possible to construct recombinant forms of various viruses, such as human immunodeficiency virus 1 (HIV-1) and hepatitis C virus (HCV), that carry foreign genes such as a reporter or marker protein in their genomes. These recombinant viruses usually faithfully mimic the life cycle of the original virus in infected cells and exhibit the same host range dependence. The development of a recombinant virus enables the efficient screening of inhibitors and the identification of specific host factors. However, to date the construction of recombinant hepatitis B virus (HBV) has been difficult because of various experimental limitations. The main limitation is the compact genome size of HBV, and a fairly strict genome size that does not exceed 1.3 genome sizes, that must be packaged into virions. Thus, the size of a foreign gene to be inserted should be smaller than 0.4 kb if no deletion of the genome DNA is to be performed. Therefore, to overcome this size limitation, the deletion of some HBV DNA is required. Here, we report the construction of recombinant HBV encoding a reporter gene to monitor the early stage of the HBV replication cycle by replacing part of the HBV core-coding region with the reporter gene by deleting part of the HBV pol coding region. Detection of recombinant HBV infection, monitored by the reporter activity, was highly sensitive and less expensive than detection using the currently available conventional methods to evaluate HBV infection. This system will be useful for a number of applications including high-throughput screening for the identification of anti-HBV inhibitors, host factors and virus-susceptible cells.

Here we describe a newly developed hepatitis B virus (HBV) reporter system to monitor the early stages of the HBV life cycle. This simplified in vitro system will aid in the screening of anti-HBV agents using a high-throughput strategy.

Desarrollo de un sistema reportero virus de la hepatitis B para supervisar las primeras etapas del ciclo de replicación

Currently, it is possible to construct recombinant forms of various viruses, such as human immunodeficiency virus 1 (HIV-1) and hepatitis C virus (HCV), ...

Fluorescence-based Neuraminidase Inhibition Assay to Assess the Susceptibility of Influenza Viruses to The Neuraminidase Inhibitor Class of Antivirals

The neuraminidase (NA) inhibitors are the only class of antivirals approved for the treatment and prophylaxis of influenza that are effective against currently circulating strains. In addition to their use in treating seasonal influenza, the NA inhibitors have been stockpiled by a number of countries for use in the event of a pandemic. It is therefore important to monitor the susceptibility of circulating influenza viruses to this class of antivirals. There are different types of assays that can be used to assess the susceptibility of influenza viruses to the NA inhibitors, but the enzyme inhibition assays using either a fluorescent substrate or a chemiluminescent substrate are the most widely used and recommended. This protocol describes the use of a fluorescence-based assay to assess influenza virus susceptibility to NA inhibitors. The assay is based on the NA enzyme cleaving the 2&#8242;-(4-Methylumbelliferyl)-&#945;-D-N-acetylneuraminic acid (MUNANA) substrate to release the fluorescent product 4-methylumbelliferone (4-MU). Therefore, the inhibitory effect of an NA inhibitor on the influenza virus NA is determined based on the concentration of the NA inhibitor that is required to reduce 50% of the NA activity, given as an IC50 value.

We describe the use of a phenotypic fluorescence-based neuraminidase inhibition assay to assess the susceptibility of influenza A and B viruses to the neuraminidase inhibitor class of antivirals.

La neuraminidasa Ensayo de inhibición basado en la fluorescencia para evaluar la susceptibilidad de los virus de la influenza a El Inhibidor de la neuraminidasa clase de antivirales

The neuraminidase (NA) inhibitors are the only class of antivirals approved for the treatment and prophylaxis of influenza that are effective against ...

Phenotypic Analysis of Rodent Malaria Parasite Asexual and Sexual Blood Stages and Mosquito Stages

Recent advances in genetics and systems biology technologies have promoted our understanding of the biology of malaria parasites on the molecular level. However, effective malaria parasite targets for vaccine and chemotherapy development are still limited. This is largely due to the unavailability of relevant and practical in vivo infection models for human Plasmodium species, most notably for P.&#160;falciparum and P. vivax. Therefore, rodent malaria species have been extensively used as practical alternative in vivo models for malaria vaccine, drug targeting, immune response, and functional characterization studies of conserved Plasmodiumspp. genes. Indeed, rodent malaria models have proven to be invaluable, especially for exploring mosquito transmission and liver stage biology, and were indispensable for immunological studies. However, there are discrepancies in the methods used to evaluate the phenotypes of transgenic and wild-type asexual and sexual blood-stage parasites.&#160;Examples of these discrepancies are the choice of an intravenous vs. intraperitoneal infection of rodents with blood-stage parasites and the evaluation of male gamete exflagellation. Herein, we detail standardized experimental methods to evaluate the phenotypes of asexual and sexual blood stages in transgenic parasites expressing reporter-gene or wild-type rodent malaria parasite species. We also detail the methods to evaluate the phenotypes of malaria parasite mosquito stages (gametes, ookinetes, oocysts, and sporozoites) inside Anopheles mosquito vectors. These methods are detailed and simplified here for the lethal and non-lethal strains of P. berghei and P. yoelii but can also be applied with some adjustments to P. chabaudi and P. vinckei rodent malaria species.

Due to the striking similarities of the life cycle and biology of rodent malaria parasites to human malaria parasites, rodent malaria models have become indispensable for malaria research. Herein, we standardized some of the most important techniques used in the phenotypic analysis of wild-type and transgenic rodent malaria species.

Análisis fenotípico de las etapas de la sangre asexual y sexual del parásito de la malaria de los roedores y las etapas de los mosquitos

Recent advances in genetics and systems biology technologies have promoted our understanding of the biology of malaria parasites on the molecular level. ...

Generating Recombinant Avian Herpesvirus Vectors with CRISPR/Cas9 Gene Editing

Herpesvirus of turkeys (HVT) is an ideal viral vector for the generation of recombinant vaccines against a number of avian diseases, such as avian influenza (AI), Newcastle disease (ND), and infectious bursal disease (IBD), using bacterial artificial chromosome (BAC) mutagenesis or conventional recombination methods. The clustered regularly interspaced palindromic repeats (CRISPR)/Cas9 system has been successfully used in many settings for gene editing, including the manipulation of several large DNA virus genomes. We have developed a rapid and efficient CRISPR/Cas9-mediated genome editing pipeline to generate recombinant HVT. To maximize the potential use of this method, we present here detailed information about the methodology of generating recombinant HVT expressing the VP2 protein of IBDV. The VP2 expression cassette is inserted into the HVT genome via an NHEJ (nonhomologous end-joining)-dependent repair pathway. A green fluorescence protein (GFP) expression cassette is first attached to the insert for easy visualization and then removed via the Cre-LoxP system. This approach offers an efficient way to introduce other viral antigens into the HVT genome for the rapid development of recombinant vaccines.

Herpesvirus of turkeys (HVT) is widely used as a vector platform for the generation of recombinant vaccines against a number of avian diseases. This article describes a simple and rapid approach for the generation of recombinant HVT-vectored vaccines using an integrated NHEJ-CRISPR/Cas9 and Cre-Lox system.

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