Abstract
Systemic application of therapeutics to the CNS tissue often results in subtherapeutic drug levels, because of restricted and selective penetration through the blood–brain barrier (BBB). Here, we give a detailed description of a standardized technique for intrathecal drug delivery in rodents, analogous to the technique used in humans. The intrathecal drug delivery method bypasses the BBB and thereby offers key advantages over oral or intravenous administration, such as maximized local drug doses with minimal systemic side effects. We describe how to deliver antibodies or drugs over several days or weeks from a s.c. minipump and a fine catheter inserted into the subdural space over the spinal cord (20 min operative time) or into the cisterna magna (10 min operative time). Drug levels can be sampled by quick and minimally invasive cerebrospinal fluid (CSF) collection from the cisterna magna (5 min procedure time). These techniques enable targeted application of any compound to the CNS for therapeutic studies in a wide range of CNS disease rodent models. Basic surgery skills are helpful for carrying out the procedures described in this protocol.
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References
Parrish, K.E., Sarkaria, J.N. & Elmquist, W.F. Improving drug delivery to primary and metastatic brain tumors: strategies to overcome the blood-brain barrier. Clin. Pharmacol. Ther. 97, 336–346 (2015).
Frank, R.T., Aboody, K.S. & Najbauer, J. Strategies for enhancing antibody delivery to the brain. Biochim. Biophys. Acta 1816, 191–198 (2011).
Pepinsky, R.B. et al. Exposure levels of anti-LINGO-1 Li81 antibody in the central nervous system and dose-efficacy relationships in rat spinal cord remyelination models after systemic administration. J. Pharmacol. Exp. Ther. 339, 519–529 (2011).
Sampson, F.C., Hayward, A., Evans, G., Morton, R. & Collett, B. Functional benefits and cost/benefit analysis of continuous intrathecal baclofen infusion for the management of severe spasticity. J. Neurosurg. 96, 1052–1057 (2002).
Gold, R. & Oreja-Guevara, C. Advances in the management of multiple sclerosis spasticity: multiple sclerosis spasticity guidelines. Expert Rev. Neurother. 13, 55–59 (2013).
Nance, P. et al. Intrathecal baclofen therapy for adults with spinal spasticity: therapeutic efficacy and effect on hospital admissions. Can. J. Neurol. Sci. 22, 22–29 (1995).
Dworkin, R.H. et al. Interventional management of neuropathic pain: NeuPSIG recommendations. Pain 154, 2249–2261 (2013).
Garg, T., Bhandari, S., Rath, G. & Goyal, A.K. Current strategies for targeted delivery of bio-active drug molecules in the treatment of brain tumor. J. Drug Target. 23, 865–887 (2015).
Bottros, M.M. & Christo, P.J. Current perspectives on intrathecal drug delivery. J. Pain Res. 7, 615–626 (2014).
Sindou, M. Neurosurgery for Spasticity: A Practical Guide for Treating Children and Adults (Springer, 2014).
Dario, A. & Tomei, G. A benefit-risk assessment of baclofen in severe spinal spasticity. Drug Saf. 27, 799–818 (2004).
Van Damme, P. & Robberecht, W. Developments in treatments for amyotrophic lateral sclerosis via intracerebroventricular or intrathecal delivery. Expert Opin. Investig. Drugs 23, 955–963 (2014).
Bonnan, M. et al. Intrathecal rituximab therapy in multiple sclerosis: review of evidence supporting the need for future trials. Curr. Drug Targets 15, 1205–1214 (2014).
Yaksh, T.L. & Rudy, T.A. Chronic catheterization of the spinal subarachnoid space. Physiol. Behav. 17, 1031–1036 (1976).
Yaksh, T.L. & Rudy, T.A. Analgesia mediated by a direct spinal action of narcotics. Science 192, 1357–1358 (1976).
LoPachin, R.M., Rudy, T.A. & Yaksh, T.L. An improved method for chronic catheterization of the rat spinal subarachnoid space. Physiol. Behav. 27, 559–561 (1981).
Yaksh, T.L. & Stevens, C.W. Simple catheter preparation for permitting bolus intrathecal administration during chronic intrathecal infusion. Pharmacol. Biochem. Behav. 25, 483–485 (1986).
Hayes, C.S., Mulkmus, S.A., Cizkova, D., Yaksh, T.L. & Hua, X.Y. A double-lumen intrathecal catheter for studies of modulation of spinal opiate tolerance. J. Neurosci. Methods 126, 165–173 (2003).
Storkson, R.V., Kjorsvik, A., Tjolsen, A. & Hole, K. Lumbar catheterization of the spinal subarachnoid space in the rat. J. Neurosci. Methods 65, 167–172 (1996).
Pogatzki, E.M., Zahn, P.K. & Brennan, T.J. Lumbar catheterization of the subarachnoid space with a 32-gauge polyurethane catheter in the rat. Eur. J. Pain 4, 111–113 (2000).
Sakura, S., Hashimoto, K., Bollen, A.W., Ciriales, R. & Drasner, K. Intrathecal catheterization in the rat. Improved technique for morphologic analysis of drug-induced injury. Anesthesiology 85, 1184–1189 (1996).
Tsang, B.K., He, Z., Ma, T., Ho, I.K. & Eichhorn, J.H. Decreased paralysis and better motor coordination with microspinal versus PE10 intrathecal catheters in pain study rats. Anesth. Analg. 84, 591–594 (1997).
van den Hoogen, R.H. & Colpaert, F.C. Long term catheterization of the lumbar epidural space in rats. Pharmacol., Biochem., Behav. 15, 515–516 (1981).
Zhao, R.R. et al. Combination treatment with anti-Nogo-A and chondroitinase ABC is more effective than single treatments at enhancing functional recovery after spinal cord injury. Eur. J. Neurosci. 38, 2946–2961 (2013).
Gonzenbach, R.R. et al. Delayed anti-nogo-a antibody application after spinal cord injury shows progressive loss of responsiveness. J. Neurotrauma 29, 567–578 (2012).
Wahl, A.S. et al. Neuronal repair. Asynchronous therapy restores motor control by rewiring of the rat corticospinal tract after stroke. Science 344, 1250–1255 (2014).
Gonzenbach, R.R. et al. Nogo-A antibodies and training reduce muscle spasms in spinal cord-injured rats. Ann. Neurol. 68, 48–57 (2010).
Malkmus, S.A. & Yaksh, T.L. Intrathecal catheterization and drug delivery in the rat. Methods Mol. Med. 99, 109–121 (2004).
Brosamle, C., Huber, A.B., Fiedler, M., Skerra, A. & Schwab, M.E. Regeneration of lesioned corticospinal tract fibers in the adult rat induced by a recombinant, humanized IN-1 antibody fragment. J. Neurosci. 20, 8061–8068 (2000).
Liebscher, T. et al. Nogo-A antibody improves regeneration and locomotion of spinal cord-injured rats. Ann. Neurol. 58, 706–719 (2005).
Boster, A.L. et al. Best practices for intrathecal baclofen therapy: dosing and long-term management. Neuromodulation. 19, 623–631 (2016).
Filli, L., Zorner, B., Weinmann, O. & Schwab, M.E. Motor deficits and recovery in rats with unilateral spinal cord hemisection mimic the Brown-Sequard syndrome. Brain 134, 2261–2273 (2011).
Yaksh, T. & Malkmus, S. Animal models of intrathecal and epidural drug delivery. Spinal Drug Deliv. 1, 317–344 (1999).
Katzung, B.G., Masters, S.B. & Trevor, A.J. Basic & Clinical Pharmacology Vol. 8 (Lange Medical Books/McGraw-Hill, New York, 2004).
Essex Wynter, W. Four cases of tubercular meningitis in which paracentesis of the theca vertebralis was performed for the relief of fluid pressure. Lancet 137, 981–982 (1891).
Shen, D.D., Artru, A.A. & Adkison, K.K. Principles and applicability of CSF sampling for the assessment of CNS drug delivery and pharmacodynamics. Adv. Drug Deliv. Rev. 56, 1825–1857 (2004).
Sakka, L., Coll, G. & Chazal, J. Anatomy and physiology of cerebrospinal fluid. Eur. Ann. Otorhinolaryngol. Head Neck Dis. 128, 309–316 (2011).
Carp, R.I., Davidson, A.L. & Merz, P.A. A method for obtaining cerebrospinal fluid from mice. Res. Vet. Sci. 12, 499 (1971).
Long, J.B., Mobley, W.C. & Holaday, J.W. Neurological dysfunction after intrathecal injection of dynorphin A (1–13) in the rat. I. Injection procedures modify pharmacological responses. J. Pharmacol. Exp. Ther. 246, 1158–1166 (1988).
Westergren, I. & Johansson, B.B. Changes in physiological parameters of rat cerebrospinal fluid during chronic sampling: evaluation of two sampling methods. Brain Res. Bull. 27, 283–286 (1991).
Pegg, C.C., He, C., Stroink, A.R., Kattner, K.A. & Wang, C.X. Technique for collection of cerebrospinal fluid from the cisterna magna in rat. J. Neurosci. Methods 187, 8–12 (2010).
Butler, M. et al. Spinal distribution and metabolism of 2′-O-(2-methoxyethyl)-modified oligonucleotides after intrathecal administration in rats. Neuroscience 131, 705–715 (2005).
Jasmin, L. & Ohara, P.T. Long-term intrathecal catheterization in the rat. J. Neurosci. Methods 110, 81–89 (2001).
Jones, L.L. & Tuszynski, M.H. Chronic intrathecal infusions after spinal cord injury cause scarring and compression. Microsc. Res. Tech. 54, 317–324 (2001).
Begley, D. et al. The role of brain extracellular fluid production and efflux mechanisms in drug transport to the brain. The Blood-Brain Barrier and Drug Delivery to the CNS, Vol. 1, 93–108 (CRC Press, 2000).
Silverberg, G.D. et al. The cerebrospinal fluid production rate is reduced in dementia of the Alzheimer's type. Neurology 57, 1763–1766 (2001).
Chiu, C. et al. Temporal course of cerebrospinal fluid dynamics and amyloid accumulation in the aging rat brain from three to thirty months. Fluids Barriers CNS 9, 3 (2012).
Ummenhofer, W.C., Arends, R.H., Shen, D.D. & Bernards, C.M. Comparative spinal distribution and clearance kinetics of intrathecally administered morphine, fentanyl, alfentanil, and sufentanil. Anesthesiology 92, 739–753 (2000).
Krupp, J.L. & Bernards, C.M. Pharmacokinetics of intrathecal oligodeoxynucleotides. Anesthesiology 100, 315–322 (2004).
Penn, R.D., Kroin, J.S., York, M.M. & Cedarbaum, J.M. Intrathecal ciliary neurotrophic factor delivery for treatment of amyotrophic lateral sclerosis (phase I trial). Neurosurgery 40, 94–99 (1997).
Weinmann, O. et al. Intrathecally infused antibodies against Nogo-A penetrate the CNS and downregulate the endogenous neurite growth inhibitor Nogo-A. Mol. Cell. Neurosci. 32, 161–173 (2006).
Zemmar, A. et al. Neutralization of Nogo-A enhances synaptic plasticity in the rodent motor cortex and improves motor learning in vivo. J. Neurosci. 34, 8685–8698 (2014).
Vuillemenot, B.R. et al. Recombinant human tripeptidyl peptidase-1 infusion to the monkey CNS: safety, pharmacokinetics, and distribution. Toxicol. Appl. Pharmacol. 277, 49–57 (2014).
Samaranch, L. et al. Strong cortical and spinal cord transduction after AAV7 and AAV9 delivery into the cerebrospinal fluid of nonhuman primates. Hum. Gene Therapy 24, 526–532 (2013).
Salegio, E.A. et al. Distribution of nanoparticles throughout the cerebral cortex of rodents and non-human primates: implications for gene and drug therapy. Front. Neuroanat. 8, 9 (2014).
Nedergaard, M. Neuroscience. Garbage truck of the brain. Science 340, 1529–1530 (2013).
Iliff, J.J. et al. A paravascular pathway facilitates CSF flow through the brain parenchyma and the clearance of interstitial solutes, including amyloid β. Sci. Transl. Med. 4, 147ra111 (2012).
Banks, W.A. The CNS as a target for peptides and peptide-based drugs. Expert Opin. Drug Deliv. 3, 707–712 (2006).
Cooper, P.R. et al. Efflux of monoclonal antibodies from rat brain by neonatal Fc receptor, FcRn. Brain Res. 1534, 13–21 (2013).
Button, K.S. et al. Power failure: why small sample size undermines the reliability of neuroscience. Nat. Rev. Neurosci. 14, 365–376 (2013).
Pardridge, W.M. Drug and gene targeting to the brain with molecular Trojan horses. Nat. Rev. Drug Discov. 1, 131–139 (2002).
Yu, Y.J. et al. Therapeutic bispecific antibodies cross the blood-brain barrier in nonhuman primates. Sci. Transl. Med. 6, 261ra154 (2014).
Vlieghe, P. & Khrestchatisky, M. Medicinal chemistry based approaches and nanotechnology-based systems to improve CNS drug targeting and delivery. Med. Res. Rev. 33, 457–516 (2013).
Tsai, S.Y., Papadopoulos, C.M., Schwab, M.E. & Kartje, G.L. Delayed anti-nogo-a therapy improves function after chronic stroke in adult rats. Stroke 42, 186–190 (2011).
Keeley, R.J., Bye, C., Trow, J. & McDonald, R.J. Strain and sex differences in brain and behaviour of adult rats: learning and memory, anxiety and volumetric estimates. Behav. Brain Res. 288, 118–131 (2015).
Jonasson, Z. Meta-analysis of sex differences in rodent models of learning and memory: a review of behavioral and biological data. Neurosci. Biobehav. Rev. 28, 811–825 (2005).
Whishaw, I.Q. & Pellis, S.M. The structure of skilled forelimb reaching in the rat: a proximally driven movement with a single distal rotatory component. Behav. Brain Res. 41, 49–59 (1990).
Montoya, C.P., Campbell-Hope, L.J., Pemberton, K.D. & Dunnett, S.B. The 'staircase test': a measure of independent forelimb reaching and grasping abilities in rats. J. Neurosci. Methods 36, 219–228 (1991).
Soblosky, J.S., Colgin, L.L., Chorney-Lane, D., Davidson, J.F. & Carey, M.E. Ladder beam and camera video recording system for evaluating forelimb and hindlimb deficits after sensorimotor cortex injury in rats. J. Neurosci. Methods 78, 75–83 (1997).
Maier, I.C. et al. Constraint-induced movement therapy in the adult rat after unilateral corticospinal tract injury. J. Neurosci. 28, 9386–9403 (2008).
Herrlich, S., Spieth, S., Messner, S. & Zengerle, R. Osmotic micropumps for drug delivery. Adv. Drug Deliv. Rev. 64, 1617–1627 (2012).
Pritchett-Corning, K. et al. Handbook of Clinical Signs in Rodents and Rabbits 1 1–136 (Charles River Laboratories, 2010).
Gelderd, J.B. & Chopin, S.F. The vertebral level of origin of spinal nerves in the rat. Anat. Rec. 188, 45–47 (1977).
Harrison, M. et al. Vertebral landmarks for the identification of spinal cord segments in the mouse. NeuroImage 68, 22–29 (2013).
Ngai, S.H., Berkowitz, B.A., Yang, J.C., Hempstead, J. & Spector, S. Pharmacokinetics of naloxone in rats and in man: basis for its potency and short duration of action. Anesthesiology 44, 398–401 (1976).
Quintela, T. et al. Sex-related differences in rat choroid plexus and cerebrospinal fluid: a cDNA microarray and proteomic analysis J. Neuroendocrinol. 28 http://dx.doi/org/10.1111/jne.12340 (2016).
Strohl, K.P. et al. Ventilation and metabolism among rat strains. J. Appl. Physiol. 82, 317–323 (1997).
Acknowledgements
This study was supported by grants from the Swiss National Science Foundation (to M.E.S.), the Christopher and Dana Reeve Foundation, the Swiss MS Society, the Hartmann-Müller Foundation, Zurich, the Desirée-and-Niels-Yde Foundation (to B.V.I.) and two MD–PhD fellowships from the Swiss National Science Foundation (no. 323530_151488, to B.V.I., and no. 323630_151489, to M.P.S.). We thank N.K. Medtner and F. Busoni for help with the surgery.
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B.V.I., L.S. and M.G. developed the operations. B.V.I., J.K. and M.P.S. performed the operations. J.K., B.V.I., N.G. and A.C.M. provided figures. J.K. designed graphics and processed videos. B.V.I., M.E.S. and M.P.S. wrote the manuscript. M.E.S., B.V.I. and M.L. contributed to conceptual development.
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Supplementary Figures 1 and 2
Supplementary Figure 1 Direct intrathecal application of antibodies and drugs to the rodent central nervous system: spinal cord. Supplementary Figure 2 Direct intrathecal application of antibodies and drugs to the rodent central nervous system: cisterna magna. (PDF 1747 kb)
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Ineichen, B., Schnell, L., Gullo, M. et al. Direct, long-term intrathecal application of therapeutics to the rodent CNS. Nat Protoc 12, 104–121 (2017). https://doi.org/10.1038/nprot.2016.151
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DOI: https://doi.org/10.1038/nprot.2016.151
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