diffusion mri

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DIFFUSION MRI SECOND EDITION


DIFFUSION MRI SECOND EDITION Edited by

HEIDI JOHANSEN-BERG AND

TIMOTHY E.J. BEHRENS Centre for Functional Magnetic Resonance Imaging of the Brain (FMRIB) Nuffield Department of Clinical Neurosciences, University of Oxford, John Radcliffe Hospital, Oxford, UK

AMSTERDAM • BOSTON • HEIDELBERG • LONDON NEW YORK • OXFORD • PARIS • SAN DIEGO SAN FRANCISCO • SINGAPORE • SYDNEY • TOKYO Academic Press is an imprint of Elsevier


Yet to come


Contents 4.4. Modified Imaging Techniques that Yield Less-Distorted

Foreword ix Contributors xi

Images 68

4.5. Imaging Techniques that Acquire Information about the OffResonance Field

I

Distortions

References 82

1. Introduction to Diffusion MR 3

5. Gaussian Modeling of the Diffusion Signal 85

¨ ZARSLAN PETER J. BASSER AND EVREN O

What is Diffusion? 3 Magnetic Resonance and Diffusion Diffusion in Neural Tissue 7 Concluding Remarks 8 Acknowledgments 9 References 9

DEREK K. JONES

5.1. 5.2. 5.3. 5.4. 5.5. 5.6.

5

2. Pulse Sequences for Diffusion-weighted MRI 11 JIM PIPE

2.1. 2.2. 2.3. 2.4. 2.5. 2.6.

MRI Pulse Sequence Primer 11 Adding Diffusion Weighting to a Pulse Sequence Bulk Motion Sensitivity 21 Single-Shot Echo Planar Imaging Methods 23 Parameter Optimization 26 Other DWI Pulse Sequences 29 References 34

88

KIRAN K. SEUNARINE AND DANIEL C. ALEXANDER

17

6.1. 6.2. 6.3. 6.4. 6.5. 6.6. 6.7.

KARLA L. MILLER

The Modular Nature of Diffusion Sequences Improving Image Quality 38 Improving Diffusion Contrast 48 Conclusions 56 Acknowledgments 57 References 57

Introduction 85 Diffusion Basics 86 Basic Modeling and Quantification Data Acquisition Strategies 97 Artifacts 98 What is a Model? 100 References 100

6. Multiple Fibers: Beyond the Diffusion Tensor 105

3. Diffusion Acquisition: Pushing the Boundaries 35 3.1. 3.2. 3.3. 3.4.

75

4.7. Recent Work at the FMRIB 77

INTRODUCTION TO DIFFUSION MRI

1.1. 1.2. 1.3. 1.4.

70

4.6. Image Registration-Based Methods for Correcting

35

Introduction 105 Multiple Fibers: What’s All the Fuss About? 106 Model-Based Approaches 108 Nonparametric Algorithms 111 Derived Information 118 Applications and Exploitation 119 Summary 120 Appendix A. Qball implementation 121 Appendix B. Spherical Deconvolution Implementation Acknowledgments 121 References 121

121

II DIFFUSION MRI FOR QUANTITATIVE MEASUREMENT

4. Geometric Distortions in Diffusion MRI 61

7. White Matter Structure: A Microscopist’s View 127

JESPER L.R. ANDERSSON

JULIA M. EDGAR AND IAN R. GRIFFITHS

4.1. Introduction 61 4.2. Echo Planar Imaging 62 4.3. Where Does the Off-Resonance Field Come From? 66

7.1. Introduction 127 7.2. Cellular Components of the CNS White Matter 130 7.3. Water Content of White Matter 145

v


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CONTENTS

7.4. Changes in White Matter Due to Abnormalities in

Myelin 145 7.5. The Ultrastructural Effects of Demyelination and Axonal Damage in Humans 147 7.6. Plasticity in White Matter 148 7.7. Summary 148 Acknowledgments 148 References 149

8. The Biological Basis of Diffusion Anisotropy 155

10.8. Statistical Modeling, Thresholding, and Multivariate Approaches

Complex Tract Structure

Microstructure 158 8.3. Role of the Apparent Diffusion Coefficients for Interpreting Anisotropy 171 8.4. Issues Related to Diffusion Anisotropy Measurements in Tissue by MRI 175 8.5. Summary 178 Acknowledgments 178 References 178

9. Inferring Microstructural Information of White Matter from Diffusion MRI 185 YANIV ASSAF AND YORAM COHEN

9.1. The Morphological Features of White Matter 185 9.2. Diffusion MRI and Tissue Microstructure 186 9.3. Diffusion Tensor ImagingdA Tool for White Matter Microstructural Mapping

187

226

10.11. Standard Space Templates and Atlases 229 10.12. Empirical Studies of Gaussianity and Repeatability in Diffusion MRI Data

230

10.13. Example Multi-Subject Studies 231 10.14. Conclusions 236 References

CHRISTIAN BEAULIEU

8.1. Utility of Microscopic Water Motion 155 8.2. Relationship of Water Diffusion Anisotropy to Tissue

219

10.9. Alternative Diffusion Measures to Test 223 10.10. Interpretation Issues: Partial Volume Effects and

236

11. Diffusion MRI in Neurological Disorders 241 BENEDETTA BODINI AND OLGA CICCARELLI

11.1. 11.2. 11.3. 11.4.

Introduction 241 Brief Overview of Methods for Clinical Research Clinical Applications 243 Conclusions 252 References 252

242

12. Diffusion Tensor Imaging in the Study of Aging and Age-associated Neural Disease 257 DAVID H. SALAT

12.1. Introduction 257 12.2. Typical Diffusion Metrics Utilized in the Study of Tissue Microstructure Across the Lifespan

258

12.3. Diffusion in Aging 261 12.4. Associations Between DTI Metrics and Gray Matter Morphometry

270

9.4. Diffusion Tensor ImagingdA Tool for White Matter

12.5. Caveats to the use of Diffusion Imaging in the Study

9.5. 9.6. 9.7. 9.8.

12.6. Future Directions 274

Microstructural Mapping? 188 Types of Diffusion Processes in the Tissue 189 Q-Space Analysis 191 Models of Diffusion in White Matter 195 Towards Virtual Biopsy of White Matter With Diffusion MRI 199 9.9. Summary 204 References 205

10. Cross-subject Comparison of Local Diffusion MRI Parameters 209 STEPHEN M. SMITH, GORDON KINDLMANN, AND SAAD JBABDI

of Aging and Age-Associated Disease

Acknowledgments References 275

271

275

13. Diffusion Imaging in the Developing Brain 283 SERENA J. COUNSELL, GARETH BALL, ANAND PANDIT, AND A. DAVID EDWARDS

13.1. Changes in Diffusion Measures with Increasing Gestational Age 283

13.2. Abnormal White Matter and Cortical Gray Matter Development in Preterm Infants at Term 285

10.1. Introduction 210 10.2. Cross-Subject Registration (Image Alignment) 210 10.3. Voxel-Based MorphometrydOverview and Application

13.3. Assessing the Connectome in the Developing

10.4. Problems of Interpretability in VBM-Style

Brain and Association with Neurodevelopmental Outcome 292 13.6. MRI in the Term Infant with Perinatal Brain Injury 294 13.7. Future Directions 296 13.8. Conclusions 297 References 297

to Diffusion Data

Analyses

212

212

10.5. Region-of-Interest and Tractography-Based Strategies for Localizing Change

214

10.6. Tract-Based Spatial Statistics 215 10.7. Other Skeleton-Based Work 218

Brain

286

13.4. DTI in Preterm Brain Injury 288 13.5. Diffusion MRI Studies of the Developing Preterm


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CONTENTS

14. Individual Differences in White Matter Microstructure in the Healthy Brain 301 JAN SCHOLZ, VALENTINA TOMASSINI, AND HEIDI JOHANSEN-BERG

14.1. Introduction 301 14.2. Gender and Handedness 303 14.3. Changes in White Matter Microstructure with Development and Aging are Associated with Development or Deterioration in Cognitive Skills 305 14.4. Age-Independent Variation in Brain Structure Reflects InterIndividual Variation in Behavior 306 14.5. Are Individual Differences in White Matter due to Nature or Nurture? 312 14.6. Conclusion 313 References 314

15. Diffusion Tensor Imaging and its Application to Schizophrenia and Related Disorders 317 M. KUBICKI, C-F. WESTIN, O. PASTERNAK, AND M.E. SHENTON

15.1. Introduction 317 15.2. Review of Dti Findings in Schizophrenia 321 15.3. Future Directions: What are we Missing and How Can we Fill in the GAPS? References 330

17.3. Contemporary Application of Experimental Tract Tracing in Non-Human Primates

17.4. Conclusions 394

References 394 Further Reading 399

18. The Human Connectome: Linking Structure and Function in the Human Brain 401 OLAF SPORNS

18.1. 18.2. 18.3. 18.4. 18.5.

What is the Connectome? 401 Modes of Brain Connectivity 402 Defining Network Nodes of the Connectome 405 Graph Analysis of Brain Connectivity 409 Mapping the Network of Structural Connections of the Human Brain 415 18.6. Relating Structural Connections to Functional Interactions 419 18.7. Brain Connectivity and Network Disease 422 18.8. The Future of the Connectome 423 Acknowledgments 423 References 423

327

16. Mapping Connections in Humans and Non-Human Primates: Aspirations and Challenges for Diffusion Imaging 335 DAVID C. VAN ESSEN, SAAD JBABDI, STAMATIOS N. SOTIROPOULOS, CHARLES CHEN, KRIKOR DIKRANIAN, TIM COALSON, JOHN HARWELL, TIMOTHY E.J. BEHRENS, AND MATTHEW F. GLASSER

16.1. 16.2. 16.3. 16.4. 16.5.

Introduction 336 Neuroanatomical Fundamentals 336 Approaches to Imaging Human Brain Connectivity 339 Imaging Structural Connectivity: The HCP Strategy 340 The Fiber Architecture of Gyral Blades and Deep White Matter 344 16.6. Discussion 352 Acknowledgments 354 References 354

365

19. MR Diffusion Tractography 429 TIMOTHY E.J. BEHRENS, STAMATIOS N. SOTIROPOULOS, AND SAAD JBABDI

19.1. 19.2. 19.3. 19.4. 19.5. 19.6.

Introduction 429 Streamline Tractography 430 Probabilistic Tractography 435 Propagation of Uncertainty in Tractography 439 Global Tractography Approaches 441 Choice of Local Description of Diffusion in Tractography 443 19.7. Designing a Diffusion Tractography Study 444 19.8. Future Advances in Diffusion Tractography 445 19.9. Statistics on Tractography 447 19.10. Summary and Conclusions 448 References 448

20. Validation of Tractography 453 PENNY L. HUBBARD AND GEOFFREY J.M. PARKER

III DIFFUSION MRI FOR IN VIVO NEUROANATOMY 17. Classic and Contemporary Neural Tract-tracing Techniques 359 ROBERT J. MORECRAFT, GABRIELLA UGOLINI, JOSE´ L. LANCIEGO, FLORIS G. WOUTERLOOD, AND DEEPAK N. PANDYA

17.1. Introduction 360 17.2. A Brief Historical Perspective of the Development of Experimental Tract Tracing

360

20.1. 20.2. 20.3. 20.4.

Introduction 453 Validation of Fiber Orientation Information Validation of Tractography 460 Summary 476 References 477

455

21. Connectivity Fingerprinting of Gray Matter 481 JOHANNES C. KLEIN, TIMOTHY E.J. BEHRENS, AND HEIDI JOHANSEN-BERG

21.1. Introduction 481 21.2. Application to Subcortical Gray Matter 488


viii

CONTENTS

21.3. Application to Cortical Gray 494 21.4. Validation 500 21.5. Conclusions 506 References

506

ˆ ME SALLET, ERIE D. BOORMAN, MATTHEW F.S. RUSHWORTH, JE´RO AND ROGIER B. MARS

22. Contribution of Diffusion Tractography to the Anatomy of Language 511 MARCO CATANI AND SANJA BUDISAVLJEVIC

22.1. Introduction 511 22.2. The Anatomy of the Arcuate Fasciculus: From Blunt Dissections to Tractography

512

22.3. Lateralization of the Arcuate Fasciculus 514 22.4. Comparative Anatomy of Perisylvian Language 22.5. 22.6. 22.7. 22.8.

24. Comparing Connections in the Brains of Humans and Other Primates Using Diffusion-weighted Imaging 569

Network 516 Functional Correlates of Perisylvian Language Network 517 Beyond the Arcuate Fasciculus: Ventral and Frontal Networks 518 Application of Tractography to Language Disorders 519 Summary and Future Directions 524 Acknowledgments 525 References 525

24.1. Introduction 569 24.2. Comparing Tractography with Tract-Tracing Techniques

570

24.3. Using Tractography to Examine the Connections of Human Ventral Frontal Cortex

571

24.4. Language and the Arcuate Fascicle in Humans and other Primates

573

24.5. Tractography Suggests Basic Similarities in Frontal Cortex Organization in Humans and other Primates

573

24.6. Premotor Cortex 578 24.7. Comparing the Parietal Cortex in Humans and other Primates

579

24.8. Conclusions 581 References

581

25. Imaging Structure and Function 585 SAAD JBABDI

23. Presurgical Tractography Applications 531 ¨ RGY A. HOMOLA ANDREAS J. BARTSCH, ARMIN BILLER, AND GYO

23.1. Introduction 531 23.2. Presurgical Applications, Tract Latitudes, and Tracking Failures

533

23.3. Potential Surgical Targets and Intentions 545 23.4. Presurgical Tractography 553 23.5. Summary and Conclusions 560 Acknowledgments References 561

561

25.1. 25.2. 25.3. 25.4. 25.5.

Introduction 585 Structural Imaging and Brain Morphometry 586 Combining Sources of Data 591 Imaging Anatomo-Functional Networks 597 Conclusions 602 References 602

Index 607


Foreword

p0010

chart pathways that relate to abilities that are unique to humans, such as language. Third, we can establish the borders between neighboring cytoarchitectonic areas in the human brain, using the principle that each area has a unique pattern of inputs and outputs. Finally, we can make use of the in vivo nature of the measurements to examine how particular white matter connections contribute to individual variability in behavior, are subject to experience-dependent plasticity, and are affected in different disease states. After introducing the key methods underlying diffu- p0055 sion imaging (Chapters 1 through 6), Section II of this book focuses on this last point: How can we make quantitative measurements of white matter connections in the living brain, and use these measurements as a probe in health and disease. In answering this question it is clearly important to know how the diffusion signal relates to the underlying biophysical properties of the axonethe real quantities of interest in white matter. Chapters 7 through 9 explore this issue in detail and examine what biological inferences we can already make from our diffusion data, and what new inferences we may be able to make in the near future. Chapters 10 through 15 then offer an essential guide to any scientist using these techniques to ask questions about white matter changes in health and disease. They start with a detailed and thorough guide to experimental design and data analysis techniques (Chapter 10), and then proceed with chapters giving key examples and highlighting important results from diffusion imaging in neurological and psychiatric disorders, in development and aging, and in behavioral neuroscience. The first two sections of this book are therefore essen- p0060 tial reading for scientists using diffusion imaging in their quantitative investigations, for those developing new methodologies for diffusion imaging, and for any clinicians and systems neuroscientists with an interest in white matter. But there is an overarching reason why we need to p0065 know about the underlying architecture of connections in the human brain. The aim of neuroscience is to understand how the brain works as a whole, and the outstanding advantage of imaging methods (fMRI, EEG, MEG) is that they are whole-brain methods. Of course, we need to understand how each area performs its specific

If you want to know how an MP3 player works, the first thing you need to establish is how it is wired up. It is the same with the brain. Yet, as Crick and Jones commented in Nature in 1993 (Crick and Jones, 1993) it was lamentable how little we knew at that time about the connections of the human brain. All we could do was infer the connections indirectly from tracer studies in non-human primates. Yet it was only a year later that Basser et al. (1994) (see Chapter 1) showed that one could use MRI to measure the diffusion of water along axons, and in this way to visualize the major fiber tracts. p0040 Anatomists were skeptical that much would come of the new methods. The reason is that their concern is with the fine details of the connections, rather than with the lie of the tracts. These fine details can be demonstrated by using the transport of tracers, and the method has been extensively used for the non-human primate brain (see Chapter 17). We currently have data for 7009 sites in the macaque brain, with 36 994 connections details (http://cocomac.g-node.org). But, unfortunately, though one can use MRI to visualize the transport of tracers (Saleem et al., 2002), it is not ethical to inject tracers into the living brain. And so far, little progress has been made in using tracers in post-mortem brains. With current methods, dyes only diffuse at the rate of about 5 mm in 2 weeks (Kobbert et al., 2000). p0045 So to what extent can diffusion MRI provide similar information for the human brain? Fortunately, methods for tractography are steadily increasing in sophistication and although challenges remain it is now possible to estimate the probabilities of connections and to trace through regions of fiber complexity with some success (Chapter 19). Confidence in the exact site of termination will increase as the spatial resolution of MRI increases and it is possible to measure anisotropy within the gray matter. There are already hopeful signs. By use of a specialized coil array it has been possible to visualize thalamocortical fibers as they penetrate perpendicular to the pial surface and terminate in layer IV of occipital cortex (Jaermann et al., 2008). And it has even been possible to measure laminar profiles of activity using fMRI (Ress et al., 2007). p0050 Given these tools, what can we use them for? First, we can check whether our inferences about connections from non-human primates are correct. Second, we can

ix


x

FOREWORD

function, and for this we will need to resort to recording the electrical activity of cells, whether singly or in subpopulations. But tasks are not performed by single areas but by systems. And this means that we have to understand the interactions between areas within those systems. In imaging, measures of effective connectivity, whether structural equation modeling or dynamic causal modeling (Penny et al., 2004), require the specification of a prior anatomical model for that system. This model need not include every synaptic stage in a circuit, but we do need to be confident that it holds for the human brain. Diffusion MRI promises to provide that model. This means that everyone who uses functional brain imaging needs to consult this book. In it they will find everything they need. p0070 The last section provides an extensive account of the uses of diffusion MRI for neuroanatomy, and it is this section that those engaged in fMRI and MEG will need to consult. It starts with an overview of the emerging science of connectomics by Van Essen et al. and Sporns (Chapters 16 and 18) and an account of the related classical methods for tract tracing in animals by Morecraft and colleagues (Chapter 17). In Chapter 19, Behrens and colleagues give an introduction to probabilistic methods for tractography. In Chapter 20 Hubbard and Parker review the ways in which tractography has been validated. Klein et al. (Chapter 21) introduce the notion of connectional fingerprints (see also Chapter 19), and demonstrate that the pattern of connections can be used to distinguish between neighboring cytoarchitectonic areas. In Chapter 22, Catani and Budisavljevic describe the language pathways in the human brain, and this chapter is followed by a discussion of the use of tractography in neurosurgical planning by

Bartsch et al. (Chapter 23). In the next chapter, Rushworth et al. compare the frontal and parietal lobe connections in the macaque and human brain (Chapter 24). The last chapter (Chapter 25) is called “Imaging structure and function.” That surely is the aim of all of those who use imaging to understand the workings of the human brain. Richard Passingham

Professor of Cognitive Neuroscience, Department of Experimental Psychology, University of Oxford, UK

References Basser, P.J., Mattiello, J., LeBihan, D., 1994. Estimation of the effective self-diffusion tensor from the NMR spin echo. J. Magn. Reson. B. 103, 247e254. Crick, F., Jones, E., 1993. Backwardness of human neuroanatomy. Nature 361, 109e110. Jaermann, T., De Zanche, N., Staempfli, P., Pruessmann, K.P., Valavanis, A., Boesiger, P., Kollias, S.S., 2008. Preliminary experience with visualization of intracortical fibers by focused highresolution diffusion tensor imaging. AJNR Am. J. Neuroradiol. 29, 146e150. Kobbert, C., Apps, R., Bechmann, I., Lanciego, J.L., Mey, J., Thanos, S., 2000. Current concepts in neuroanatomical tracing. Prog. Neurobiol. 62, 327e351. Penny, W.D., Stephan, K.E., Mechelli, A., Friston, K.J., 2004. Modelling functional integration: a comparison of structural equation and dynamic causal models. NeuroImage 23 (Suppl. 1), S264eS274. Ress, D., Glover, G.H., Liu, J., Wandell, B., 2007. Laminar profiles of functional activity in the human brain. NeuroImage 34, 74e84. Saleem, K.S., Pauls, J.M., Augath, M., Trinath, T., Prause, B.A., Hashikawa, T., Logothetis, N.K., 2002. Magnetic resonance imaging of neuronal connections in the macaque monkey. Neuron 34, 685e700.


Contributors Daniel C. Alexander Department of Computer Science, University College London, UK

Olga Ciccarelli NMR Unit, Queen Square MS Centre, UCL Institute of Neurology, London, UK

Jesper L.R. Andersson Centre for Functional Magnetic Resonance Imaging of the Brain (FMRIB), Nuffield Department of Clinical Neurosciences, University of Oxford, John Radcliffe Hospital, Oxford, UK

Tim Coalson Department of Anatomy and Neurobiology, Washington University School of Medicine, St. Louis, MO, USA Yoram Cohen School of Chemistry, Raymond and Beverly Sackler Faculty of Exact Sciences, Tel Aviv University, Tel Aviv, Israel

Yaniv Assaf Department of Neurobiology, George S. Wise Faculty of Life Sciences, Tel Aviv University, Tel Aviv, Israel

Serena J. Counsell Centre for the Developing Brain, Department of Perinatal Imaging, Division of Imaging Sciences & Biomedical Engineering, St Thomas’ Hospital Kings College London, London, UK

Gareth Ball Centre for the Developing Brain, Department of Perinatal Imaging, Division of Imaging Sciences & Biomedical Engineering, St Thomas’ Hospital Kings College London, London, UK

A. David Edwards Centre for the Developing Brain, Department of Perinatal Imaging, Division of Imaging Sciences & Biomedical Engineering, St Thomas’ Hospital Kings College London, London, UK

Andreas J. Bartsch Department of Neuroradiology, University of Heidelberg, Heidelberg, Germany; Centre for Functional Magnetic Resonance Imaging of the Brain (FMRIB), Nuffield Department of Clinical Neurosciences, University of Oxford, John Radcliffe Hospital, Oxford, UK

Krikor Dikranian Department of Anatomy and Neurobiology, Washington University School of Medicine, St. Louis, MO, USA

Peter J. Basser Section on Tissue Biophysics and Biomimetics, PPITS, NICHD, National Institutes of Health, Bethesda, MD, USA

Julia M. Edgar Institute of Infection, Immunity and Inflammation, College of Medical, Veterinary and Life Sciences, University of Glasgow, Glasgow, UK; Department of Neurogenetics, Max Planck Institute of Experimental Medicine, Goettingen, Germany

Christian Beaulieu Department of Biomedical Engineering, Faculty of Medicine and Dentistry, University of Alberta, Canada

Matthew F. Glasser Department of Anatomy and Neurobiology, Washington University School of Medicine, St. Louis, MO, USA

Timothy E.J. Behrens Centre for Functional Magnetic Resonance Imaging of the Brain (FMRIB), Nuffield Department of Clinical Neurosciences, University of Oxford, John Radcliffe Hospital, Oxford, UK; Department of Experimental Psychology, University of Oxford, Oxford, UK

Ian R. Griffiths Institute of Infection, Immunity and Inflammation, College of Medical, Veterinary and Life Sciences, University of Glasgow, Glasgow, UK

Armin Biller Department of Neuroradiology, University of Heidelberg, Heidelberg, Germany

John Harwell Department of Anatomy and Neurobiology, Washington University School of Medicine, St. Louis, MO, USA

Benedetta Bodini Centre de recherche de l’institut du cerveau et de la moelle e´pinie`re, Universite´ Pierre & Marie Curie, Hoˆpital Pitie´ Salpeˆtrie`re, Paris, France

Gyo¨rgy A. Homola Department of Neuroradiology, University of Wu¨rzburg, Wu¨rzburg, Germany

Erie D. Boorman Department of Experimental Psychology, University of Oxford and Centre for Functional Magnetic Resonance Imaging of the Brain (FMRIB), Nuffield Department of Clinical Neurosciences, University of Oxford, John Radcliffe Hospital, Oxford, UK

Penny L. Hubbard Centre for Imaging Sciences & Biomedical Imaging Institute, University of Manchester, Manchester Academic Health Science Centre, Manchester, UK Saad Jbabdi Centre for Functional Magnetic Resonance Imaging of the Brain (FMRIB), Nuffield Department of Clinical Neurosciences, University of Oxford, John Radcliffe Hospital, Oxford, UK

Sanja Budisavljevic Natbrainlab, Section of Brain Maturation and Centre for Neuroimaging Sciences, Institute of Psychiatry, King’s College London, UK

Heidi Johansen-Berg Centre for Functional Magnetic Resonance Imaging of the Brain (FMRIB), Nuffield Department of Clinical Neurosciences, University of Oxford, John Radcliffe Hospital, Oxford, UK

Marco Catani Natbrainlab, Section of Brain Maturation and Centre for Neuroimaging Sciences, Institute of Psychiatry, King’s College London, UK Charles Chen Department of Anatomy and Neurobiology, Washington University School of Medicine, St. Louis, MO, USA

Derek K. Jones Cardiff University Brain Research Imaging Centre (CUBRIC), School of Psychology, Cardiff University, UK

xi


xii

CONTRIBUTORS

Gordon Kindlmann Department of Computer Science, University of Chicago, Chicago, USA Johannes C. Klein Department of Neurology, GoetheUniversity of Frankfurt, Frankfurt am Main, Germany; Centre for Functional Magnetic Resonance Imaging of the Brain (FMRIB), Nuffield Department of Clinical Neurosciences, University of Oxford, John Radcliffe Hospital, Oxford, UK M. Kubicki Psychiatry Neuroimaging Laboratory, Department of Psychiatry, Brigham and Women’s Hospital, Harvard Medical School, Boston, MA, USA Jose´ L. Lanciego Center for Applied Medical Research (CIMA and CIBERNED), Neurosciences Division, University of Navarra Medical College, Pamplona Navarra, Spain Rogier B. Mars Department of Experimental Psychology, University of Oxford and Centre for Functional Magnetic Resonance Imaging of the Brain (FMRIB), Nuffield Department of Clinical Neurosciences, University of Oxford, John Radcliffe Hospital, Oxford, UK Karla L. Miller Centre for Functional Magnetic Resonance Imaging of the Brain (FMRIB), Nuffield Department of Clinical Neurosciences, University of Oxford, John Radcliffe Hospital, Oxford, UK Robert J. Morecraft Division of Basic Biomedical Sciences, Laboratory of Neurological Sciences, University of South Dakota, Sanford School of Medicine, Vermillion, South Dakota, USA ¨ zarslan Section on Tissue Biophysics and BiomiEvren O metics, PPITS, NICHD, National Institutes of Health, Bethesda, MD, USA Anand Pandit Centre for the Developing Brain, Department of Perinatal Imaging, Division of Imaging Sciences & Biomedical Engineering, St Thomas’ Hospital Kings College London, London, UK Deepak N. Pandya Department of Anatomy and Neurobiology, Department of Neurology, Boston University School of Medicine, Boston, Massachusetts, USA Geoffrey J.M. Parker Centre for Imaging Sciences & Biomedical Imaging Institute, University of Manchester, Manchester Academic Health Science Centre, Manchester, UK O. Pasternak Psychiatry Neuroimaging Laboratory, Department of Psychiatry, Brigham and Women’s Hospital, Harvard Medical School, Boston, MA, USA; Laboratory of Mathematics in Imaging, Department of Radiology, Brigham and Women’s Hospital, Harvard Medical School, Boston, MA, USA Jim Pipe Keller Center for Imaging Innovation, Barrow Neurological Institute, Phoenix, Arizona, USA

Matthew F.S. Rushworth Department of Experimental Psychology, University of Oxford and Centre for Functional Magnetic Resonance Imaging of the Brain (FMRIB), Nuffield Department of Clinical Neurosciences, University of Oxford, John Radcliffe Hospital, Oxford, UK David H. Salat Athinoula A. Martinos Center for Biomedical Imaging, Charlestown, MA, USA, Massachusetts General Hospital Department of Radiology, Boston, MA, USA; Boston VA Healthcare System, Neuroimaging Research for Veterans Center, Boston, MA, USA Je´roˆme Sallet Department of Experimental Psychology, University of Oxford and Centre for Functional Magnetic Resonance Imaging of the Brain (FMRIB), Nuffield Department of Clinical Neurosciences, University of Oxford, John Radcliffe Hospital, Oxford, UK Jan Scholz Hospital for Sick Children, Toronto, Ontario, Canada Kiran K. Seunarine Department of Computer Science, University College London, UK M.E. Shenton Psychiatry Neuroimaging Laboratory, Department of Psychiatry, Brigham and Women’s Hospital, Harvard Medical School, Boston, MA, USA; VA Boston Healthcare System, Brockton, MA, USA Stephen M. Smith Centre for Functional Magnetic Resonance Imaging of the Brain (FMRIB), Nuffield Department of Clinical Neurosciences, University of Oxford, John Radcliffe Hospital, Oxford, UK Stamatios N. Sotiropoulos Centre for Functional Magnetic Resonance Imaging of the Brain (FMRIB), Nuffield Department of Clinical Neurosciences, University of Oxford, John Radcliffe Hospital, Oxford, UK Olaf Sporns Department of Psychological and Brain Sciences, Indiana University, Bloomington, IN, USA Valentina Tomassini Institute of Psychological Medicine and Clinical Neurosciences, Cardiff University School of Medicine, University Hospital of Wales, Heath Park, Cardiff, UK Gabriella Ugolini Laboratoire de Neurobiologie Cellulaire et Molećulaire (NBCM), CNRS, Gif-sur-Yvette, France David C. Van Essen Department of Anatomy and Neurobiology, Washington University School of Medicine, St. Louis, MO, USA C.-F. Westin Laboratory of Mathematics in Imaging, Department of Radiology, Brighann and Women’s Hospital, Harvard Medical School, Boston, MA, USA Floris G. Wouterlood Department of Anatomy and Neurosciences, Vrije Universiteit Medical Center, Neuroscience Campus Amsterdam, Amsterdam, The Netherlands

S E C T I O N

I I I

DIFFUSION MRI FOR IN VIVO NEUROANATOMY

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Chapter: 17

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C H A P T E R

17 Classic and Contemporary Neural Tract-tracing Techniques

c0017

Robert J. Morecraft*, Gabriella Ugoliniy, Jose´ L. Lanciegoz, Floris G. Wouterloodx, Deepak N. Pandyaxx *

Division of Basic Biomedical Sciences, Laboratory of Neurological Sciences, University of South Dakota, Sanford School of Medicine, Vermillion, South Dakota, USA, yLaboratoire de Neurobiologie Cellulaire et Molećulaire (NBCM), CNRS, Gif-sur-Yvette, France, zCenter for Applied Medical Research (CIMA and CIBERNED), Neurosciences Division, University of Navarra Medical College, Pamplona Navarra, Spain, xDepartment of Anatomy and Neurosciences, Vrije Universiteit Medical Center, Neuroscience Campus Amsterdam, Amsterdam, The Netherlands, xxDepartment of Anatomy and Neurobiology, Department of Neurology, Boston University School of Medicine, Boston, Massachusetts, USA

O U T L I N E 17.1 Introduction

17.3.2.5 Final Stages of Tissue Processing 17.3.3 Specific Tract-Tracing Techniques 17.3.3.1 Horseradish Peroxidase (HRP) 17.3.3.2 Lectins (Wheat Germ Agglutinin) and Toxins (Cholera Toxin B Fragment) 17.3.3.3 Fluorescent Tracing Compounds 17.3.3.4 Dextrans 17.3.3.5 Phaseolus vulgaris Leucoagglutinin (PHA-L) 17.3.3.6 Combined Neuroanatomical Tract-Tracing Methods 17.3.4 Transneuronal Tracers 17.3.4.1 Transneuronal Tracing Using Conventional Tracers 17.3.4.2 Viral Transneuronal Tracers

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17.2 A Brief Historical Perspective of the Development of Experimental Tract Tracing 360 17.2.1 Early Period of Gross Dissection 360 17.2.2 Early Microscopic Observations, Enhanced Tissue Preparation and the Development of Histological Applications 361 17.2.3 Experimental Tract Tracing by Means of Neural Degeneration 364 17.3 Contemporary Application of Experimental Tract Tracing in Non-Human Primates 365 17.3.1 Basic Mechanisms Underlying Intra-Axonal Transport of Tract Tracers 365 17.3.2 The Basic Anatomical Tract-Tracing Procedure 369 17.3.2.1 Administration of Tract Tracer 369 17.3.2.2 Post-Injection Survival Period 371 17.3.2.3 Tissue Fixation and Sectioning 371 17.3.2.4 Detection of the Transported Tract Tracer 372

Diffusion MRI http://dx.doi.org/10.1016/B978-0-12-396460-1.00017-2

372 373 373 373 374 376 377 378 386 386 387

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17.1 INTRODUCTION

For centuries, investigation of the organization of nerve pathways has been of fundamental interest in neurology and the neurosciences. Indeed, how the nervous system is structured and how this organization relates to function in healthy as well as diseased states has propelled this curiosity. It is without question that gradual improvements in gross dissection techniques, steady advancements in tissue preservation and histological application, accompanied by increased measures of microscopic resolution have all contributed in their own way. Consequently, these efforts have served to catalyze significant advancements in experimental methodology while simultaneously advancing our understanding of the structure and function of the nervous system. The enduring lineage of discovery has led to our present, advanced technological standing in which a previously unimaginable arsenal of tracttracing methods is available to investigate the structural and functional organization of neural pathways in experimental animals. p0015 Since there are so many new and powerful methods that can be employed in an extensive number of creative combinations, it is not possible to cover all in a single chapter. Therefore, the primary intent of this chapter is to highlight the most common tract-tracing methods currently employed in the non-human primate model. First, the reason for this focus is that numerous white matter pathways of the non-human primate brain have homologous counterparts in the human brain which are currently being studied at a feverish pace with in vivo diffusion MRI. Second, major functional subdivisions of the non-human primate cerebral cortex have been shown to have homologous counterparts in the human brain, including, to some degree, multimodal association cortex. Related to this is that non-human primates have relatively well-developed cognitive and behavioral capacities. Third, observations from nonhuman primate studies currently serve as the “gold standard” upon which many of the in vivo experiments conducted in the human model are designed and evaluated. Indeed, in contrast to the older methods, we can now with razor-sharp precision differentiate the transported tracing compounds from competing levels of background artifact. Finally, the level at which nerve pathways are currently being investigated in the human brain draws upon critical non-human primate experimental data that has been gathered with both classical as well as contemporary tract-tracing methodologies. p0020 To put our modern fiber tracing capabilities into a historical perspective, we will draw attention to some landmark events that have contributed to the scientific evolution of neuron tract tracing. Indeed, technological

advancement built upon these monumental achievements continues to this day to be the driving force in tract-tracing discovery. The closing stages of this chapter are devoted to providing a glimpse at some highly technical and eloquent methodologies that are currently being used to define functional, cellular, molecular, and spatial features of neural pathways in non-human primates. We will conclude by listing some important references authored by experts in the field of experimental tract tracing (Further Reading). To the interested reader, these will provide more detailed information about these tract-tracing methods, including the laboratory procedures required for their implementation in experimental animals and more in-depth considerations of their specific advantages, disadvantages, and limitations than can be covered in our chapter.

17.2 A BRIEF HISTORICAL PERSPECTIVE s0010 OF THE DEVELOPMENT OF EXPERIMENTAL TRACT TRACING 17.2.1 Early Period of Gross Dissection

s0015

The earliest tract-tracing approaches employed gross p0025 dissection as the exclusive tool of exploration (Finger, 1994; Clark and O’Malley, 1996; Swanson, 2000; Schmahmann and Pandya, 2006). As limited as this technique was, because it relies heavily on the fine visuomotor skills of the dissector, it paved the way for the discovery of hundreds of fascinating features of CNS organization. The Greek physician Galen (AD 130–200) was among the first on record to dissect animals and provide detailed documentation of his observations. Many of his greatest works depict the nervous system and some of his most influential theories concerned the function of the brain. Galen believed that the ventricles contained the “vital spirits,” a blood-derived medium produced in the heart, which was stored in the ventricles and transported on demand through the “hollow” peripheral nerves to subserve motor and sensory function. His reliance on animal models for drawing these conclusions was of necessity, since dissection of the human body was forbidden by the ruling Roman Empire. Galen’s theories had a dominant influence on the medical profession and, surprisingly, lasted without challenge for more than 1000 years. But as soon as authoritative restrictions on human dissection were relaxed and “autopsies,” a term coined by Galen, became a matter of routine, many of Galen’s structural observations and functional interpretations were inevitably re-evaluated and redefined. Perhaps the most renowned anatomist to revolu- p0030 tionize our understanding of the nervous system after Galen was the Belgian physician Andreas Vesalius

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(1514–1564), who in 1543 published his classic anatomical masterpiece entitled De Humani Corporis Fabrica (On the Structure of the Human Body) (Figure 17.1) (Finger, 1994). His work was conducted in the Renaissance cities of Padua and Venice where he was provided with a generous supply of human material for dissection. Vesalius’ published figures continue to be viewed to this day, captivating interest and being admired by medical professionals as well as the lay public (Figures 17.1 and 17.2). Indeed, Vesalius and his artistic assistants of speculative identity set a new standard of excellence in the art of medical illustration (Zimber, 2001). He was the first to clearly differentiate the superficially located gray matter of the cerebrum from the harder, white substance located below (Figure 17.2, right) (Finger, 1994; Swanson, 2000; Schmahmann and Pandya, 2006).

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He also noted the structural continuity established between the underlying white matter and the corpus callosum and recognized its stable relationship with the opposite hemisphere (Figure 17.2, left). Vesalius documented numerous central and peripheral nervous system structures. Among the central white matter structures he distinguished the corona radiata, internal capsule, external capsule, and cerebellar peduncles (Figure 17.2, right). After discovering numerous faults in Galen’s work he encouraged his contemporaries to launch a new era of discovery, intended to challenge the doctrines that had ruled medicine for more than 1000 years (Singer, 1952; Swanson, 2000).

17.2.2 Early Microscopic Observations, Enhanced Tissue Preparation and the Development of Histological Applications

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At the end of the sixteenth century, the Dutch p0035 spectacle-makers Hans and Zacharias Jansen built the first compound microscope. This imaginative optical instrument opened new vistas in scientific exploration in the field of tracing of nerve tracts in the CNS. However, because of the primitive nature of the device, dissection

Two figures of dorsal view of the brain at various f0015 stages of dissection from Andreas Vesalius’ Fabrica. Left: Dorsal view of the cerebral hemispheres with the calvaria removed and dura matter reflected. The right and left hemispheres are gently retracted laterally, exposing the medial wall of the hemisphere and the corpus callosum (L) located in the depths of the interhemispheric fissure. The falx cerebri and the attached inferior sagittal sinus are resting on the dorsolateral convexity of the left hemisphere. Right: Partial axial view of the anterior half of the cerebral hemisphere and some anatomical components in the region of the posterior cranial fossa including the tentorium cerebelli (O) covering most of the cerebellum. In the cerebral hemispheres, the subcortical interface between the cortical gray matter and underlying bed of white matter is clearly demarcated. Also beautifully depicted in the region of the subcortical white matter are the corona radiata, internal capsule (E) and external capsule. Distinguished subcortical gray matter structures include the caudate nucleus, putamen, globus pallidus and thalamus. Vesalius did not name these structures and it is thought that his intent to localize them in his figures was simply to emphasize the distinction between gray and white matter entities.

FIGURE 17.2

f0010 FIGURE 17.1

The magnificent title page from Andreas Vesalius’ Fabrica. The complete title, Andreae Vesalii Bruxellensis, Scholae medicorum Patauinae professoris, de Humani corporis fabrica Libri septem, is translated as “Andreas Vesalius of Brussels, professor at the school of medicine at Padua, on the fabric of the Human body in seven books”. In the view of many historical authorities, this text ranks as one of the most important books in medical history.


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remained the primary tool for CNS investigation, while in the background, technical developments in the design of the microscope slowly carried on. In 1664 the Italian Marcello Malpighi, regarded by some as the founder of microscopic anatomy and the first histologist, boiled nervous tissue in water and provided the first microscopic description of white matter. Notably, his observations suggested nerve tissue was composed of “fibers.” Significant improvements in the microscope were made by the English physicist Robert Hooke (1635–1703) who was the first to designate the term “cell” from a biological perspective. He selected this term because of the compartmentalized appearance of plant cells which reminded him of the architectural configuration of monks’ quarters which were at the time referred to as “cellula.” The Dutchman Antonius van Leeuwenhoek (1632–1723) became one of the most renowned scientists of the seventeenth century because he possessed genuine, unabated scientific inquiry and a unique skill to fabricate the best microscope lenses of the time (Finger, 1994). He reportedly possessed hundreds

of self-made microscopes and, in a letter sent to the Royal Society of London in 1674, provided the first detailed account of the microscopic anatomy of a nerve fiber (Figure 17.3) (Clark and O’Malley, 1996). Subsequent advances in tissue preparation included p0040 boiling nervous tissue in oil (Raymond de Vieussens, 1635–1715), hardening the brain in alcohol (Felix Vicq d’Azyr, 1748–1794), and adding potash or ammonia to alcohol (Christian Reil, 1759–1813). All these innovations supported the advancement of finer, more detailed dissections of the human brain, revealing more and more of the three-dimensional organization of this remarkable organ (Figure 17.4) (Swanson, 2000; Schmahmann and Pandya, 2006). Nicolaus Steno’s (Niels Stensen, 1638–1686) scraping method of dissection, designed to follow or trace fiber bundles from one definable end to the other, led to significant advancements in our understanding of the anatomical organization of the CNS in the seventeenth and eighteenth centuries (Schmahmann and Pandya, 2006). Undeniably, gross dissection of the CNS and anatomical discovery continued to flourish, and the

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17.3 Schematic representation of Antonius van Leeuwenhoek’s microscopic observation of the cross-section of the optic nerve redrawn from his original work (van Leeuwenhoek, 1719). In the center of the figure is a small funiculus with individually drawn nerve fibers. The perineurial sheaths of adjacent funiculi without the nerve fibers surround the central funiculus. In the central funiculus the axon fibers appear squeezed and in slit-like form surrounded by the myelin sheath which appears to be swollen (Ochs, 1979).

FIGURE 17.4 Reproduction of a dissection of the ventral view of f0025

the cerebral cortex from Fe´lix Vicq d’Azyr’s atlas of the human brain (1786). Note the exquisite three-dimensional details of the hippocampus and lateral ventricle. Reproduced with permission of Dr. Larry Swanson, University of Southern California.


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synthesis of these structural observations led to an abundance of theories regarding the functional organization of the brain (Finger, 1994). p0045 In the 1800s several important developments paved the way for more powerful methods to trace fibers in the nervous system. Foremost among these were innovative advances in histological laboratory techniques and improved microscope optics (Finger, 1994; Swanson, 2000; Schmahmann and Pandya, 2006). The most significant laboratory advances were improved tissue fixation through chromic acid and formaldehyde, the development of the microtome for precision serial tissue sectioning, and the widespread application of staining nerve tissue with dyes. In 1862 the German pathologist Carl Weigert developed the first myelin stain. Formulating the mathematical principles of microscope lenses, the German physicist Ernst Abbe (1840–1905) designed lenses with corrected spherical and chromatic aberrations which significantly improved the optical performance of the microscope. Together, the advances in histological preparation and improved resolution provided by the microscope offered a new dimension in tracing major fiber pathways in the brain. Influential figures in this field included Paul Emil Flechsig (1847–1929), who used the myelin stain to describe the developing patterns of fiber myelination (Van Hoesen, 1993), Carl Wernicke (1848–1900), who in addition to contributing to the field of cortical localization published several atlases of the human brain based on myelin-stained material (Schmahmann and Pandya, 2006), and Joseph Jules Dejerine (1849–1917), a French neurologist who described numerous clinical syndromes and used myelin-stained preparations to illustrate the organization of association fiber systems in great detail in his classical work Anatomie des centres nerveux, published in 1895 (Bassetti and Jagella, 2006; Schmahmann and Pandya, 2006). However, it was challenging to trace small, complex fiber pathways in myelin-stained serial sections since fibers could be lost through densely packed plexi, or they could curve within the section, confounding and even losing fiber continuity with adjacent tissue sections (Nauta, 1993). This remained a major problem even with the discovery of stains that could selectively label axon fibers, such as the Bielschowsky method published in 1902. Furthermore, the myelin stains were unpredictable when detecting poorly myelinated fibers and, obviously, unable to identify non-myelinated fiber bundles. p0050 The existence of nerve fibers was an accepted concept before the discovery of the neuron soma. However, the recognition of both as nervous tissue constituents triggered a debate whether or not these were actually attached entities. Several investigators, including Robert Remak (1815–1865) and Jan Evangelista Purkinje (1787–1869), reported that nerve cells indeed had

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branches or extensions. In 1865 Otto Friedrich Karl Deiters (1834–1863) and his colleague Max Schultz (1825–1874) described the somal appendages of the neuron as the axis cylinder (later termed “axon” by Albert von Ko¨lliker in 1896) and protoplasmic processes (finally termed “dendrites” by Wilhelm His in 1889) (Finger, 1994; Bentivoglio, 1996). Camillo Golgi’s (1843–1926) silver-staining technique, published in 1873, put an end to the debate as it exquisitely demonstrated the neuron soma and its attached processes. Golgi referred to his method as the reazione nera (“black reaction”) as it beautifully stained neuronal cell bodies along with their dendritic processes and axons. His technique is still in use today, particularly in post-mortem human work and developmental studies (Rosoklija et al., 2003; Friedland et al., 2006). Like all methods, Golgi’s reazione nera had its own p0055 shortcomings which continue to confront modern-day investigators: random and inconsistent staining of neuron subpopulations, poor affinity to myelin, and limited use in tracing axons over long distances in serial sections. However, its application in the hands of Golgi’s rival, the famous Spanish neuroanatomist Santiago Ramon y Cajal (1852–1934), paved the way for a myriad of brilliant descriptions of the nervous tissue milieu. In this epoch when scientists conceived the nervous system still as a syncytium, Ramon y Cajal became the first and foremost advocate of the neuron theory. His observations provided support for the novel concept of nerve contacts (Foster, 1897) and he went on to describe many important elements of cortical, cerebellar, and spinal neuronal architecture. He also overcame the difficulty in tracing axon pathways over long distances with the Golgi method by studying these processes in animals with small brains (Glickstein, 2006). As translated in Clark and O’Malley (1996), Ramon y Cajal eloquently summarized his observations on the general plan of nervous tissue organization in his 1894 Croonian lecture by stating: In a synthetic manner one can say that the whole nerve center is the result of association of the four following paths: the nerve cells with short axis cylinders, that is, branching in the very thickness of the gray matter; the terminal nerve fibers which come from other centers or distant regions of the same center; nerve cells with a long axis cylinder, that is, extending as far as the white matter; the collaterals which originate either during the passage of axis cylinder extensions of the cells with long processes [axons] across the gray matter, or during the course of the tubes [bundles] of white matter.

In 1906 Golgi and Ramon y Cajal shared the Nobel p0060 Prize for Medicine because of their pioneering efforts in advancing our understanding of the microscopic organization of the nervous system (Glickstein, 2006). The Golgi method and its variants (Golgi–Cox, Golgi– Colonnier, and many others) have recently been


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f0030 FIGURE 17.5 Golgi–Cox impregnated layer III pyramidal neuron of the mouse brain, visualized by traditional brightfield microscopy (a) as

well as under the confocal microscope (b and c). By using the so-called “reflection method,” a flattened projection of the stained neuron is generated from the Z stack, therefore leading to complete visualization of the dendritic arborization. Furthermore, by taking advantage of laser zooming, small neuronal processes such as thin dendrites and dendritic spines became clearly visible.

revitalized, since despite some limitations, these methods still represent the best way to visualize small neuronal processes such as dendritic spines. Indeed, visualizing Golgi-stained neurons under the confocal microscopy has been made possible by using the socalled “reflection method” (Figure 17.5), a procedure that maximizes the detection of the laser emission reflected from stained specimen, such as the case for metallic stains (Spiga et al., 2011; Lanciego and Wouterlood, 2011).

s0025 17.2.3 Experimental Tract Tracing by Means of Neural Degeneration p0065

In the second half of the nineteenth century many investigators such as Ludwig Tu¨rck, Bernhard von Gudden, Constantin von Monakow, and David Ferrier began to study anterogradely degenerating fiber pathways in serial tissue sections following brain lesions in experimental animals (Swanson, 2000; Schmahmann and Pandya, 2006). This clever approach to tracing fiber systems was applied following Augustus Waller’s classic observation in 1850, which demonstrated that part of the nerve fiber located distal to the injury site progressively degeneratesda process that is still referred to as Wallerian degeneration (Waller, 1850). This observation also suggested to Waller that cell bodies located upstream of the lesion site may provide a nutritional and trophic influence on the attached fiber. In the 1880s Carl Weigert took advantage of Wallerian degeneration in combination with his myelin stain to infer pathway direction by microscopic observation of fiber system atrophy or cavitation. A parallel and complementary approach to the study of nerve interconnections was applied in 1879 by Von Gudden who recognized retrograde somal degeneration of cell bodies in the brainstem after severing cranial nerves as they emerge from the cranial vault. This, in combination with Franz Nissl’s pioneering application of aniline dyes, which he used

to accurately define the various stages of chromatolysis in degenerating cell bodies following axon injury, clearly indicated that the degeneration method had great potential to study both the origin of nerve pathways, as well as their point of termination. A major breakthrough in the field of tractography p0070 emerged in 1885 when the Italian histologist Marchi and his colleague Algeri published their classical method of tracing axons by following the decomposing myelin sheath as a consequence of an experimentally induced brain injury (Waller, 1850; Marchi and Algeri, 1885; Manni and Troiani, 2005). In the Marchi technique, the myelinated fibers were severed from their origin by a localized lesion of the gray matter. This trauma was followed by a predetermined survival period and subsequently by tissue fixation to capture the progressive degeneration of the myelin. Importantly, this method permitted the tracing of microscopically verified fiber pathways over long distances. In the early to midtwentieth century, the Marchi technique was the primary method used to trace axonal connections in experimental animals, resulting in a growing body of literature on cortical and subcortical connectivity in the non-human primate brain (Mettler, 1935a, 1935b, 1935c, 1935d; Kreig, 1954). Importantly, this also provided a critical source of neuroanatomical information for interpreting neurological complications and syndromes as well as developing neurosurgical treatments for the relief of involuntary movement disorders (Meyers, 1951; Walker, 1952). A significant improvement in the experimental p0075 approach to tracing CNS pathways arrived when subsequent to a lesion, the degenerating “axon” was visualized in tissue preparations 66 years after the first publication by Marchi. The need to identify the degenerating axon and not the myelin sheath for fiber tracing was brilliantly recognized by the late Walle Nauta (1916–1994), who, as a student at the university of Leiden, the oldest University in the Netherlands, noted that sparsely myelinated and unmyelinated axons in the hypothalamic region of the


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rat brain were undetectable using Marchi’s myelin staining technique. Another important issue was the potential identification of the end ramifications of pathways and the synaptic terminals at their very ends. To overcome these limitations, Nauta toiled in the laboratory with silver stains for more than 20 years, interrupted intermittently by his education and the German occupation of his homeland during World War II (Nauta, 1993). A major breakthrough occurred by virtue of a fortunate “accident,” in which by necessity he used old formalin to prepare his reducer for one of the experimental reactions. Nauta stated in reference to the developed tissue sections: “when I examined them under the microscope I saw, for the first time in all those years, a picture of axon degeneration that resembled the one I had been hoping for” (Nauta, 1993). p0080 It was recognized by his colleague Paul Gygax that the stale formalin had high concentrations of formic acid and that it had been the combination of fresh formaldehyde acidified with appropriate levels of formic acid that had stained degenerating axons black (Nauta and Gygax, 1951). A major problem encountered with this method was that both normal axons as well as degenerating axons became staineddbut it did form a new starting point. To address this issue, Nauta and Gygax devised a “suppressive” silver method which minimized the staining of healthy or normal fibers while retaining the robust visualization of the degenerating axons (Nauta and Gygax, 1954) (Figure 17.6). They also noted that their method in some cases could identify patchy areas of what appeared to be degenerating fiber terminals. The introduction of the suppressive silver method led to an escalation of neuroanatomical tracttracing studies outlining important corticocortical (e.g. Myers, 1962; Nauta, 1964; Kuypers et al., 1965; Pandya and Kuypers, 1969; Jones and Powell, 1970; Van Hoesen et al., 1972), corticolimbic (Nauta, 1972), and corticofugal (e.g. Johnson et al., 1968; Kuypers, 1981) relationships in the non-human primate brain and parallel advances in neurological interpretations of this information. p0085 The final modification in Nauta’s axon degeneration technique occurred in the mid-1960s at the Massachusetts Institute of Technology when Lennart Heimer, in collaboration with Robert Fink, made improvements which permitted the consistent visualization of degenerating terminals (Figure 17.7) (Fink and Heimer, 1967). As intuitively noted by Nauta, in spite of the promise of the new Fink–Heimer method, this improvement was destined to “compose a brief chapter in the 90-yearlong history of tracing fibers by means of experimentally induced Wallerian degeneration” (Nauta, 1993). As will be described below, a new wave of tract-tracing methods was on the immediate horizon that capitalized on the in vivo uptake and subsequent intra-axonal transport of compounds injected directly into intact nervous tissue.

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The important commonality of the new and powerful p0090 neuroanatomical tract tracers developed in the late 1960s and early 1970s was that they capitalized on cytoplasmic flow mediated specifically by the axoplasmic transport system (terms in bold are defined in Box 17.1). The movement of fluid through the nerves was not a contemporary idea by any stretch of the imagination. As noted earlier in this chapter, in the second century Galen proposed that each peripheral nerve served as a “hollow channel” through which “animal spirits” move between organ systems and the ventricular core, thereby establishing a physiological route of communication between peripheral structures and the central nervous system. Likewise, at the end of the seventeenth century, van Leeuwenhoek (Figure 17.3) noted that if pressure was applied to the severed end of a nerve a clear substance would discharge from the cut surface (Clark and O’Malley, 1996). This simple but perceptive observation paved the way for numerous theories, proposed over several centuries, on the potential functions of this fluid and its role in nerve action (Ochs, 1979, 2004). However, it was not until 1948 that axoplasmic “flow,” or “streaming” was experimentally demonstrated by Weiss and Hiscoe in their classic paper entitled “Experiments on the mechanism of nerve growth.” In addition to exploiting the axonal transport system, p0095 another major advantage of the new tract-tracing approach was that damage to the nervous tissue was limited to the degree of injury inflicted by a finetipped needle, or the tapered end of a glass micropipette. In fact, some compounds could be applied to the tissue surface without injury, but the primary mode of tract tracer delivery was, and continues to be, by injection. Once the needle or micropipette is carefully lowered into the appropriate locus in the brain, the tract-tracing compound can be introduced with great precision. This approach keeps the brain largely intact compared to the extensive damage necessarily inflicted by the lesion-degenerating techniques. Simultaneously it created a new dimension for high-resolution investigation of neural interconnections and their spatial relationships. A tremendous amount of information on how p0100 injected tract-tracing compounds are processed, packaged, and transported by neurons has been obtained from experimental studies conducted with two tracers: horseradish peroxidase (HRP) (Kristensson and Olsson, 1971; LaVail and LaVail, 1972) and radiolabeled amino


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f0035 FIGURE 17.6 Photomicrographs from Pandya and Kuypers (1969) showing examples of silver impregnated fiber degeneration in the non-

human primate using the Nauta–Gygax method (Nauta and Gygax 1951) in the: periarcuate cortex (a) and precentral gyrus; (b) following a lesion of the middle and superior tempopral gyri, stria Lancisi; (c) following a a lesion of the temporal pole; intraparietal sulcus; (d) following a lesion of the periarcuate sulcus, periarcuate cortex; (e) following a lesion of the occipital lobe; precentral gyrus; (f) following a lesion of the inferior parietal lobe; precentral gyrus; (g) following a lesion of the postcentral gyrus; postcentral gyrus; (h) following a lesion of the precentral gyrus and; periarcuate cortex; (i) following a lesion of the inferior parietal lobule. Reproduced from Pandya and Kuypers (1969) with permission from Elsevier.

acids (Cowan et al., 1972; Droz, 1979; Edwards and Hendrickson, 1981). In terms of uptake, HRP internalization occurs via fluid endocytosis at either the axon terminal, neuron soma, or neuron dendrite (Figure 17.8). Once inside the neuron, endosomes (i.e. membranebound vesicles containing the tracer substance) are

formed. Endosomes typically take on an oval or tubular shape and are then attached to the axoplasmic transport system. Eventually, the HRP is transported towards lysosomes in the cell soma where it is degraded. Tritium-labeled amino acids appear to be taken up by endocytosis by the dendrites or cell bodies and


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f0040 FIGURE 17.7

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Photomicrographs from Pandya and colleagues (1971) illustrating terminal fiber labeling using the Fink–Heimer (Fink and Heimer, 1967) method which was a simple modification of the original Nauta–Gygax (Nauta and Gygax 1951) silver impregnation protocol. The panels show degenerating terminals in the monkey following callosal section in: (a) primary motor cortex; (b) primary somatosensory cortex; (c) inferior occipital sulcus; (d) second somatosensory area; (e) superior temporal gyrus; (f) superior temporal sulcus; (g) arcuate sulcus; and (h) inferior parietal lobule. Reproduced from Pandya et al. (1971) with permission from Elsevier.


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BOX 17.1 Antibodies: Proteins found in blood or other bodily fluids that are used by the immune system to localize and neutralize foreign substances, such as bacteria and viruses. ATP: Adenosine-50 -triphosphate is a nucleotide that mediates intracellular transport of chemical energy within cells for metabolism. Axoplasm: The cytoplasm in the axon of a neuron. Catabolism: Process of breaking down molecules into smaller units. Cytoplasm: Semi-transparent fluid that forms much of the volume of the cell. The cytoplasm has three major elements; the cytosol, organelles and inclusions. Cytoskeleton: Cellular scaffolding within the cytoplasm that maintains cell structure and shape, and plays an important role in intracellular transport of vesicles and organelles. There are three main types of cytoskeletal filaments: microfilaments, intermediate filaments, and microtubules. Dyneins: Class of motor proteins that move along microtubules that are powered by ATP. The active movement of kinesins assists in axon transport typically directed toward the cell body (i.e. in the retrograde direction). Endoplasmic reticulum: Cytoplasmic organelle formed by an interconnected network of tubules,

subsequently incorporated into proteins in the metabolic machinery of the endoplasmic reticulum of the cell soma. These proteins are then transported through the axon transport system towards the terminal boutons. Although the axoplasmic transport system functions independently from the electrical conduction system, the rate of compound uptake and transport rates are related to synaptic activity and can either be facilitated by electrical stimulation (Warr et al., 1981; Bilgen et al., 2006) or inhibited by systemic pentobarbital (Turner, 1977) or by experimental removal of excitatory inputs (Singer et al., 1977). The axoplasmic transport system is also energy dependent and requires calcium and ATP from nearby mitochondria. p0105 Neuronal tract tracers may preferentially utilize different features of the axon transport system. One major feature is the directionality of axoplasmic flow and another is the transport speed, or rate at which the compound moves through the axoplasm. In terms of directionality, transport from the soma to the axon terminal is referred to as anterograde transport (also known as orthograde transport). Anterograde transport

vesicles, and cisternae that fulfil specialized functions such as protein construction and calcium uptake. Immunohistochemistry: Process of localizing molecules in cells of tissue sections by employing the principle of antibody/antigen interaction. Iontophoresis: Method of propelling high concentrations of a charged substance by electromotive force using an electrical charge. Kinesins: Class of motor proteins that move along microtubules that are powered by ATP. The active movement of kinesins assists in axon transport typically directed away from the cell body (i.e. in the anterograde direction). Lysosomes: Cytoplasmic organelles that contain enzymes which digest or breakdown macromolecules (such as injected neuronal tract tracer) and old organelles. Microtubules: Hollow cylinders formed by linear polymers of the protein tubulin. They assist in the intracellular movement of vesicles and organelles. Mitochondria: Membrane-enclosed cytoplasmic organelles that have many roles, including the generation of most of the cell’s ATP that is used as a vital source of chemical energy.

is commonly used for the translocation of organelles, vesicles, macromolecules, enzymes, and neurotransmitters where its primary function seems to be to replenish membranes and proteins of the axon, to assist in axon growth, and to replenish synaptic vesicles and other components of the presynaptic axon terminals. The rate or speed of anterograde transport is generally classified as either fast or slow. The slow component travel rate is 0.1–20 mm/day, while the fast rate is in the order of 100–450 mm/day (Edwards and Hendrickson, 1981; Oztas, 2003). From a neuroanatomical perspective, anterograde transport allows for the identification of an efferent projection from a group of cell bodies involved in the injection site. Thus, anterograde tract tracers label axons together with their terminal projection fields. A classic example of a tract tracer that is transported anterogradely from the soma to the synaptic terminal and along the slow transport system is tritiumlabeled amino acids (Figure 17.9). Transport in the opposite direction, that is, from the p0110 neuron terminal to the neuron soma, is referred to as retrograde transport. Compounds transported in the


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Incoporation of HRP D. by cell body Tubules & cisterns of agranular teticulum

Multivesicular body Lysosome 1° Golgi complex

Lysosome 2°

Nissl body

Agranular reticulum

C.

Tubule of axonal Agranular reticulum

Anterograde Axonal Transport of HRP

B.

Retrograde axonal transport of HRP

Tubule or cistern of Agranular reticulum

Multivesicular body Synaptic vesicles

Various pinocytotic organelles

A. Incoporation of HRP by axon terminal f0045 FIGURE 17.8 Schematic representation of HRP endocytosis, intracellular packaging and axoplasmic transport. Redrawn from Heimer and RoBards (1981), p. 213, with permission from Springer.

retrograde direction include proteins for building the cytoskeleton such as microtubules, organelles including mitochondria, enzymes and materials taken up by endocytosis. The fast component of retrograde transport operates at a rate of 100–200 mm/day. From a neuroanatomical standpoint, retrograde transport defines the cells of origin which project to the location of the injection site. In other words, retrograde tracers label the axon and its parent cell body and in some cases portions of the dendrites and axon collaterals. Horseradish peroxidase (HRP) is a classic example of a compound transported retrogradely and along the fast transport system (Figure 17.10). However, HRP is transported along the fast anterograde system as well. Therefore reciprocity of a neural projection system is determined in this case using the same tracer and there are many other examples of contemporary tract tracers transported bidirectionally. In fact, only radioactivity built into proteins following application of specific tritiated amino acids (e.g. proline) moves exclusively anterogradely.

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Anterograde and retrograde transport is maintained p0115 and guided by the intra-axonal microtubule system, which also forms a critical component of the cytoskeleton (Brown, 2003; Bryantseva and Hoerber, 2012; Jeppesen and Hoerber, 2012). Anterograde transport is primarily mediated by kinesin family proteins. With the microtubule system forming a stable scaffold, the anterogradely transported vesicles and their contents are pulled along the microtubule lattice by movable bridges formed by kinesin. The vesicles are often referred to as “cargo” and the kinesin molecules forming the bridges between the microtubule and membrane vesicles are commonly referred to as “molecular motors.” Characteristic rates of movement are affected by the affinity of membrane-bound molecules with kinesin. Higher affinity promotes faster transportation rates. Retrograde transport, on the other hand, is mediated by the proteins of the dynein family. Like kinesin, dynein forms a molecular bridge between the transport vesicle and microtubules, but in contrast to kinesin it transports the vesicle from terminal end of the axon toward the soma. Although kinesin and dynein are generally classified as mediating anterograde and retrograde transport, respectively, recent evidence indicates that dynein and kinesin may be involved in some levels of bidirectional transport as well (Brown, 2003).

17.3.2 The Basic Anatomical Tract-Tracing Procedure

s0040

It is without question that the vast number of tract- p0120 tracing compounds available supports many different and highly creative approaches to explore the organization and connectivity of the CNS. However, when considering the classic methods as a starting point, a common sequence of procedural events forms the backbone of a typical tract-tracing experiment. The classical method requires neurosurgical exposure to inject the compound, a specified survival period to allow for the uptake and transport of the compound, appropriate fixation of the nervous tissue once the survival period is concluded, tissue processing by which the location of the tracing compound is reported by a visible product, and finalizing stages of histochemical preparation such as counterstaining and coverslipping. 17.3.2.1 Administration of Tract Tracer

s0045

Each tracer compound is commonly injected into a p0125 predetermined location of the CNS using a controlled delivery device which serves to increase target accuracy and enhance the control over the amount of tracer volume injected into the target tissue. In non-human primates commonly a Hamilton microsyringe is used.


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(a)

(b)

(c)

(d)

f0050 FIGURE 17.9

Photomicrographs illustrating an example of and injection of tritiated amino acids into the arm representation of the primary motor cortex (M1) in the rhesus monkey. (a) Brightfield image of a Nissl-stained section showing the injection site (black precipitatedsee arrow). (b) Brightfield Nissl-stained image showing the location of labeling (black box) through the upper region of the internal capsule (ic) illustrated in panel c. (c) Darkfield image of fiber labeling in the internal capsule (white arrows) and terminal labeling (white arrow heads) in the dorsomedial region of the ventral anterior nucleus of the thalamus (Th). (d) Crescent shaped field of labeled axon terminals (white arrow heads) in the dorsal region of the putamen (Pu). ca, caudate nucleus; cgs, cingulate sulcus; cs, central sulcus; GP, globus pallidus; Pu, putamen; RN, reticular nucleus. Scale bar in c applies to d.

The syringe is placed in a microinjector unit which in turn is linked to a micromanipulator. The microinjector unit allows for small increments of pressure to be applied to the plunger of the syringe which in turn results in controlled delivery of small amounts of tracer (0.1 mL–1.0 mL). The size or spread of the injection depends on multiple factors. These include needle

diameter, rate of injection, composition of the tissue, injected concentration, total volume delivered, survival time and efficacy of the perfusion and fixation procedure. The micromanipulator permits control in the x-, y-, and z-axes over the placement of the tip and is necessary for stereotaxic navigation to subcortical targets. Another common injection method utilizes a glass


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(a)

(b)

371

(c)

f0055 FIGURE 17.10

Photomicrographs showing an example of an injection of equal parts of HRP and HRP–WGA into the gyrus rectus of the orbital frontal cortex in the rhesus monkey (from case 2 of Morecraft et al., 1992). (a) Brightfield image of a neutral red stained section through the injection site (black precipitate) involving area 14. Examples of retrogradely labeled neurons in the dysgranular region of the temporal pole (TPdg) (b) and the central superior nucleus (Csn) and medial dorsal nucleus (MD) of the thalamus (c). The black arrowheads denote a patch of dense anterograde labeling. Arrowheads in panel c denote anterograde labeling. mos, medial orbital sulcus; ros, rostral sulcus. Scale bars: (a) 1 mm, (b) 200 mm, (c) 200 mm.

micropipette to deliver the tract-tracing compound. With a precision glass micropipette puller, the taper and tip diameter at the working end of the pipette can be tailored for making very small and precisely localized injections. The compound can be injected from the micropipette using iontophoresis with the aid of a constant current source, with air pressure from a pneumatic picopump, or via a hydraulic delivery system. A relatively new development is the peri- or juxtacellular injection technique, which is the delivery of a very small volume of tracer in the extracellular space immediately next to a neuron’s cell body via a micropipette used to record neurophysiological activity from that cell (Pinault, 1996; Duque and Zaborszky, 2006). Via this approach, single cells or small clusters of cells take up the tracer and become labeled in a Golgi-like fashion, including the entire axon trajectory and terminal ramification, down to the exquisite display of synaptic boutons. s0050 17.3.2.2 Post-Injection Survival Period p0130 Following injection, the tract-tracing compound must be allowed sufficient time for uptake and axoplasmic transport. The appropriate duration of the survival period after injection must be determined for each compound and the specific projection system under investigation. As discussed earlier, some tracers take advantage of the rapid axoplasmic transport system which experimentally translates into a short postinjection survival period. Horseradish peroxidase is a classic example of a neural tract tracer that is transported rapidly in both the retrograde and anterograde directions (Figure 17.10). Optimally detectable HRP reaction products are obtained following a survival

period ranging between 2 and 5 days post injection (Mesulam, 1978; Dum and Strick, 1996). On the other hand, tritiated amino acids are incorporated into cytoplasmic proteins and transported by means of the slow axon transport system (Figure 17.9). Thus, depending on the length of the fiber system of interest, optimal visualization of transport with the tritiated amino acid tracers occurs following a 7–21 day post-injection survival period in non-human primates. As will be discussed later, substances like dextran amines, however, involve even longer post-injection survival periods in the non-human primate which can extend up to several months (Alipour et al., 2002; Lacroix et al., 2004; Rosenzweig et al., 2010). 17.3.2.3 Tissue Fixation and Sectioning s0055 The primary goal of the tissue-processing procedure is p0135 to maximize the microscopic visibility of the tract-tracing compound while minimizing background levels of artifact. This process is initiated by deeply anesthetizing the animal and perfusing it with a solution of saline, or phosphate-buffered saline, to clear the circulatory system of endogenous fluids. The saline solution is subsequently followed by the infusion of a fixative, which typically is a buffered aldehyde compound such as formaldehyde or glutaraldehyde. In neuroanatomical experiments the primary purpose of fixation is to “stabilize” or “fix” the tract-tracing compound to the precise intracellular location to which it was transported and limit its diffusion from this location during subsequent processing. The fixative of choice is determined by the specific tracttracing compound employed as well as by additional procedures included in the entire tissue-processing plan. Such additional procedures may include multiple


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steps to visualize additional pathways in conjunction with the primary pathway (i.e. convergence of pathways onto a common target nucleus) or terminal affiliated neurotransmitters or receptors. These are referred to as double- or triple-labeling methods and are commonly accomplished using contemporary immunohistochemical detection methods. Multilabeling procedures take advantage of antibodies directed against the first, second, or third compound of interest which are then visualized using different colorization approaches as will be illustrated later in this chapter. Consideration of electron microscopic evaluation for synaptic contacts also affects the selection of a particular fixative. p0140 Depending on the experimental design of the tracttracing paradigm, a cryoprotectant may be infused after the fixative. Cryoprotection is required to prevent tissue degradation by the formation of large ice crystals when the tissue is frozen prior to long-term storage or cutting. Typical cryoprotectants are sucrose (in phosphate buffer) and buffered glycerin or buffered dimethyl sulfoxide/ glycerin mixtures. Such mixtures provide excellent tissue preservation and less shrinkage than sucrose (Rosene et al., 1986). Once fixed and cryoprotected the tissue is frozen and sectioned using a microtome. p0145 Alternatively, infusion of a cryoprotectant may be avoided for certain experimental designs. One example would be when the experimental design calls for postmortem intracellular filling of isolated retrogradely labeled somas to enhance visualization of the dendritic processes (McNeal et al., 2008) or when synaptic connectivity is visualized linking a series of neural projections (Jorritsma-Byham et al., 1994; Kajiwara et al., 2008). In these cases, tissue blocks are attached to a substrate and sliced wet using a vibrating microtome. Following sectioning with either the microtome or vibratome, the tissue slices are rigorously kept in serial order so the labeled pathways can be reconstructed in three dimensions with specialized microscopic applications and computer software, or alternatively, reconstructed in two dimensions as classically depicted in line drawings of serial sections or cortical surface plottings. s0060 17.3.2.4 Detection of the Transported Tract Tracer p0150 Inherent fluorescent tract-tracing compounds, fluorescent microbeads, or commercially available compounds with an attached fluorochrome marker need not be processed further after microtome sectioning if the purpose is single tracer microscopic evaluation. In these cases, the tissue sections can simply be mounted on glass slides, coverslipped, and viewed in the microscope under epifluorescence illumination. However, if the tract tracer is in an invisible state after sectioning there are a number of approaches one can take to locate the tract tracer by a visibly detectable reaction product. One of these is through histochemical processing such

as that applied to visualize HRP (Figure 17.10), or autoradiography which is used to visualize radiolabeled amino acids incorporated in neurons (Figure 17.9). Another more commonly applied method to visualize tract tracers is through immunohistochemical processing which utilizes the antibody labeling method. Briefly, immunohistochemical visualization can be achieved using a variety of methodological approaches including the use of primary and secondary antibodies, the peroxidase–anti-peroxidase (PAP) method, the avidinlabeling method, or the highly sensitive avidin–biotin– peroxidase complex (ABC) approach (Kobbert et al., 2000). The immunohistochemical method allows for considerable amplification of the tract tracer signal which leads to a highly accurate localization of labeled products. In both histochemical and immunohistochemical methods, optical visualization is ultimately achieved by the accumulation of an insoluble dense precipitate located directly over the transported compound. The latter occurs as a consequence of incubating the tissue sections in a solution containing a chromogen (colorizing agent) followed by an enzymatic reaction initiated by the presence of hydrogen peroxide. Typical chromogens are benzidine derivatives such as diaminobenzidine (DAB) and tetramethylbenzidine (TMB). More exotic chromogens include benzidine dihydrochloride (BDHC), p-phenylendiamine (PPD-PC), 1-naphthol/azur B (Mauro et al., 1985), and the magenta chromogen, Vector-VIP (Zhou and Grofova, 1995). Recently, a green substrate named HistoGreen has been made available (Thomas and Lemmer, 2005). Depending on the method applied, the resultant precipitate can be black, blue, purple, green, or brown. As the contrast of the brown diaminobenzidine reaction product is often low against the background, the addition of a trace of nickel ammonium sulfate to the reaction mixture produces a deep blue to black reaction product that significantly enhances the contrast. A sequence of a nickel-enhanced diaminobenzidine p0155 reaction followed by a “classical” diaminobenzidine reaction leads in a double-tracing experiment to differently colored precipitates (i.e. black and brown, respectively). Finally, several investigators have used alkaline phosphatase reaction (purple reaction product) as substrate instead of diaminobenzidine, or in double-label experiments in combination with diaminobenzidine (Wouterlood et al., 1987). An alternative to the dense reaction product in the immunohistochemical method is attaching a fluorochrome at the end stages of the tissue processing procedure in place of the benzidine derivative. 17.3.2.5 Final Stages of Tissue Processing s0065 In the final stages of tissue processing, the individual p0160 tissue slices are mounted on gelatin-coated glass


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microscope slides, dried, and coverslipped. Prior to coverslipping, the tissue sections may be counterstained to facilitate the cytoarchitectural details of the neural tissue in direct relation to the injection site and transported tracer. A variety of counterstains can be applied to the tissue sections for either light or fluorescence microscopic analysis. It is important that the color of the counterstain does not obstruct the detection of the reaction products for either light level or fluorescence microscopic analysis. An alternative approach is to counterstain an unreacted series of tissue sections that are immediately adjacent to the reacted series.

s0070 17.3.3 Specific Tract-Tracing Techniques s0075 17.3.3.1 Horseradish Peroxidase (HRP)

p0165

A classical approach employing a retrograde tracing method to study connectivity of the CNS is via the injection of a 20–50% solution of HRP in saline through a Hamilton microsyringe. Alternative application forms of HRP have been used which include HRP-solution drenched gel foam, crystals, pellets, and plain gel. Although occasionally being used in cortical connectional studies, the latter is primarily used for tracing peripheral nervous system pathways where typically a spinal or cranial nerve is transected and the HRP medium is applied directly to the cut end of the proximal nerve trunk. The most common type of HRP used is Sigma Type VI (Sigma-Aldrich, St Louis, MO, USA) or Boehringer (Boehringer-Ingelheim). Mechanical injections typically include volumes of 0.1–0.5 mL per injection site. Large injections must be carried out carefully to avoid mechanical damage to the brain tissue at the syringe tip which may lead to local necrosis. For this reason, administration must be slow, over a 10–20 min period. After the injection is completed the syringe is left in situ for a short period of time to avoid backfilling of the syringe track with HRP. p0170 As metabolic reduction of the enzymatic activity of transported HRP is a function of time, and transport is related to distance and typically differs between fiber systems, the optimal survival period following injection of HRP varies from 2 to 5 days. Longer transport periods usually result in poor detectability since the tracer is catabolized. The classical detection method of transported HRP is a straightforward enzyme histochemical reaction. The fixative includes a mixture of glutaraldehyde (0.5–1.5%) with 4% buffered formaldehyde (Mesulam, 1982). Tissue sections obtained by cutting the brain on a freezing microtome are first bleached with 1–2% hydrogen peroxide to suppress endogenous peroxidase activity, and then incubated for 30 min in diaminobenzidine-Tris-HCl with peroxide added. When sufficient precipitate has

373

formed, typically a brown punctate staining of the cytoplasm of neuronal cell bodies against a clear background, the reaction is terminated by transferring the sections to rinsing solution. The sections are then mounted on gelatin-coated glass slides, dried, counterstained, and coverslipped. 17.3.3.2 Lectins (Wheat Germ Agglutinin) and Toxins (Cholera Toxin B Fragment)

s0080

In the continuing search for reliable and sensitive p0175 retrograde tracers, some lectins and toxins were introduced in the 1980s as bidirectional tracers. First, a lectin, wheat germ agglutinin (WGA), was introduced as a highly sensitive retrogradely transported tracer by Schwab et al. (1978). In these initial studies, WGA was radiolabeled, but it was soon discovered that WGA could be conjugated with HRP (Schwab et al., 1978; Gonatas et al., 1979), which made it possible to detect the transported tracer using the same enzymatic reactions and substrates as for native HRP (see section above). During the following years, the transport capabilities of a large number of HRP conjugated lectins and toxins were examined and the HRP conjugates of WGA (WGA–HRP) and of the non-toxic B fragment of cholera toxin (choleragenoid) (CTB–HRP) were found to be the most sensitive, both as anterogradely and retrogradely transported tracers (Trojanowski et al., 1981, 1982). Undamaged axons of passage have only a limited capacity to internalize these probes. These conjugates are quantitatively more sensitive retrogradely and anterogradely transported markers than native HRP, since, for example, their retrograde transport labels a greater number of neurons and reveals their dendritic arborization better than native HRP. The superior sensitivity of WGA–HRP and CTB–HRP is especially due to the fact that their uptake is receptor mediated (i.e. it reflects binding of WGA and CTB to specific receptors on neuronal membranes, which is not affected by their conjugation with HRP; Trojanowski, 1983). Such binding results in uptake of the WGA–HRP and CTB–HRP conjugates by adsorptive endocytosis, a process that is more efficient than the uptake of native HRP by fluid phase endocytosis. Another fact that contributes to their greater sensitivity is that WGA–HRP and CTB– HRP are less rapidly eliminated from retrogradely labeled neurons than native HRP (Wan et al., 1982), probably because they are transported along intracellular pathways different than those utilized by native HRP (Trojanowski et al., 1982; Trojanowski, 1983). Both WGA–HRP and CTB–HRP are transported bidirectionally, and this may be an advantage or a disadvantage depending on the system under investigation. Anterograde axonal transport of WGA– HRP occurs at a rate of approximately 108 mm/day, resulting in a slower arrival of this probe at axon


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terminals compared with native HRP, which is anterogradely transported at a rate of 288–432 mm/day (Trojanowski, 1983). p0180 Recently, WGA and CTB have also been used as tracers in their unconjugated form (Luppi et al., 1990), since they can be visualized via an immunohistochemical procedure using specific primary antibodies. While peroxidase is an intrinsic marker since this enzyme is found in some form in all neurons (peroxisomes, lysosomes) and as a consequence of this may give rise to considerable background staining, a lectin such as WGA or a toxin like CTB is exotic to the brain and therefore can be visualized razor-sharp, with little to no background when detected immunohistochemically. Today, most CTB applications include either immunoperoxidase or immunofluorescence detection, via an intermediary step of incubation with an antibody against CTB, although direct tracing with CTB–rhodamine and CTB–fluorescein fluorescent conjugates has been also reported (e.g. in the visual system of mice; Lyckman et al., 2005). Indeed, conjugates of CTB with a wide variety of Ò Alexa fluorochromes have recently become available (Invitrogen-Molecular Probes). These compounds are directly visible in a fluorescence microscope and, because of their photostability they are particularly well suited for inspection in the confocal laser scanning microscope. p0185 As with any methodology there are both advantages and limitations to the application of these markers in neuroanatomical studies. The nature of the experiments planned will determine which tracers and methods are most suitable in a given situation. Some of the potential drawbacks of their use are the same as those associated with other tracers. The visualization of the injection site produced by WGA–HRP and CTB–HRP appears similar to that produced by native HRP. The extent of the apparent injection site depends upon the sensitivity of the substrate used for histochemical visualization of HRP activity (e.g. the injection site appears smaller when detected with DAB than with the more sensitive TMB substrate). Determining the effective area of uptake within the injection sites is a common problem with all these markers (Mesulam, 1982). Differences have been observed in the efficiency of uptake in given pathways, which are related to differences in availability of receptors for WGA or CTB on neuronal membranes. This is particularly the case in the peripheral nervous system, where important differences have been observed in the uptake and anterograde transganglionic transport of WGA–HRP and CTB–HRP in primary sensory neurons. For instance, a preferential uptake occurs by small primary sensory neurons for WGA–HRP and by large primary sensory neurons in the case of CTB–HRP (Robertson and Arvidsson, 1985; Robertson and Grant, 1985).

17.3.3.3 Fluorescent Tracing Compounds

s0085

Kuypers and collaborators (Kuypers et al., 1977, 1979; p0190 Bentivoglio et al., 1979) were the first to demonstrate that fluorescent dyes injected focally into the brain “behaved” similarly to HRP, that is, they are taken up by nerve terminals and transported retrogradely to the cell bodies of the involved projection neurons (Figure 17.10). Within a few years, Kuypers and collaborators introduced a great number of fluorescent tracers including Evans Blue, Bisbenzimide, Dapi-primuline, Propidium Iodide (Kuypers et al., 1979), True Blue, Granular Blue (Bentivoglio et al., 1979), Nuclear Yellow, Fast Blue (Kuypers et al., 1977, 1979, 1980; Bentivoglio et al., 1980), and Diamidino Yellow (Keizer et al., 1983). A common characteristic of these dyes is that they bind to adenine–thymine-rich nucleic acids, and can be seen directly under epifluorescence illumination, which eliminates the need for histochemical treatment. Because of their different spectral characteristics, such fluorescent markers serve as the ideal tracers to study axon collateralization in double-labeling experiments and, in combination with immunofluorescence, to study the neurochemical identity of traced fiber projections (Kuypers and Huisman, 1984). After a brief epoch of intense experimentation with p0195 numerous fluorescent compounds in non-human primates, the neuroscience community settled with the combination of Fast Blue (FB) (Bentivoglio et al., 1980) and Diamidino Yellow (DY) (Keizer et al., 1983) for retrograde double labeling (Figure 17.11c) (Kuypers and Huisman, 1984). The introduction of fluorescence retrograde double-labeling methods by Kuypers and collaborators made it finally possible to clarify neuroanatomically the extent of collateralization of axonal projections. For example, use of these techniques revealed the existence of important differences in the collateralization of descending motor pathways and made it possible to quantify them very precisely (Huisman et al., 1982; Keizer and Kuypers, 1989). In addition, the capacity of FB and DY to persist in neurons over extended periods of time has made them very powerful tools for studying the developmental remodeling of neuronal pathways (Innocenti et al., 1986; Meissirel et al., 1991). The great advantages of FB and DY compared to p0200 earlier compounds are their fixation compatibility and their similar transport time, making it possible to apply them in the same experiment (Figure 17.11c). The fact that they do not easily diffuse out of retrogradely labeled neurons also permits a wider range of survival times and less possibility of spurious labeling (Bentivoglio et al., 1980; Keizer et al., 1983; Kuypers and Huisman, 1984). Since FB and DY label different features of the cell at the same excitation wavelength (the cytoplasm


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(a)

(b)

Tpt

SII

(c)

375

Lateral Basal Nucleus

(d)

Area 28

f0060 FIGURE 17.11

Photomicrographs illustrating neurons labeled with the fluorescent tracers Fast Blue (FB) and Diamidino Yellow (DY) following their injection into the CNS of the rhesus monkey. (a) DY-labeled cells in layers III and V of the second somatosensory cortex (SII) following injection of this tracer into the face region of the primary somatosensory cortex (SI). Inset: higher magnification of the retrogradely DY-labeled neurons. Scale bar ¼ 300 mm (from Morecraft et al., 2004). (b) FB-labeled neurons in layer V of area Tpt following an injection of this tracer into the caudal region of the caudal cingulate motor cortex (M4 or cCMC). Inset: higher magnification of the retrogradely FB-labeled cells. Scale bar ¼ 300 mm (from Morecraft et al., 2004). (c) Cells in the lateral basal nucleus of the amygdala labeled retrogradely with both DY and FB (double arrowhead) following an injection of FB into the rostral cingulate motor cortex (M3 or rCMC) and DY into M4. Also in the microscopic field are cells labeled only with FB (blue cytoplasm) and only with DY (single arrowhead). Inset shows a double-labeled cell (from Morecraft et al., 2007a). (d) Extensive intermingling of FB- and DY-labeled cells in entorhinal cortex (area 28) following an injection of FB into the rostral part of M3 and an injection of DY into the middle part of M3 (from Morecraft et al., 2012). (a–c) Reproduced from Moorcraft et al. (2004, 2007b) with permission from John Wiley & Sons, Inc. (d) Reproduced from Moorcraft et al. (2012) with permission from Elsevier.

in blue for FB, and the nucleus in yellow for DY), double-labeled neurons can be easily distinguished (Figure 17.11c). Furthermore, it is also possible to use these tracers in combination with a third fluorescent marker for multiple labeling (Godschalk et al., 1984). FB can also produce intense anterograde and retrograde labeling of axons, including axon collaterals (Rosina, 1982; Ugolini and Kuypers, 1986). Axons and terminals labeled by the anterograde and retrograde transport of FB show a different labeling intensity. For instance, anterograde fiber labeling of axons is “sharp,” as compared to “soft” retrograde fiber labeling (Ugolini and Kuypers, 1986). Moreover, the labeling of axons and terminals produced by FB can be combined with retrograde labeling of the cell bodies of their target neurons by means of DY (or another fluorescent tracer). Using this labeling combination, for example, Ugolini and Kuypers (1986) were able to demonstrate that corticospinal axons send a massive number of collateral projections to pontocerebellar neurons that innervate the anterior lobe. p0205 To this day, FB and DY remain the gold standards for double retrograde fluorescent labeling of multiple

neuronal populations over long distances in primates (e.g. Keizer and Kuypers, 1989; He et al., 1993; Morecraft et al., 2004, 2007b, 2012). Application of these tracers is via mechanical injection, although micro-iontophoretical delivery of these fluorescent dyes has also been reported elsewhere. Most fluorescent tracers can be dissolved in water. However, DY is more difficult to dissolve and is typically suspended in phosphate buffer (Keizer et al., 1983) and sonicated just before use. DY produces smaller injection sites than FB, which may be an advantage depending on the system that is being investigated. Transport times for long distances in primates are usually between 1 and 5 weeks, depending on the distance (e.g. 5 weeks for strong labeling of corticospinal and corticobulbar neurons in macaque monkeys after injection of DY in the cervical spinal cord and FB in the lower medulla) (Keizer and Kuypers, 1989). The tissue is commonly fixed by perfusion fixation with phosphatebuffered formalin. Higher concentrations of formalin reportedly enhance FB fiber labeling (Ugolini and Kuypers, 1986). The tissue is then cryoprotected and cut on a freezing microtome. Sections are immediately mounted on glass slides to avoid in vitro diffusion of


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the tracer from labeled neurons (Keizer et al., 1983). The sections are then air-dried and stored in the dark at 4 C. Coverslipping requires a special non-fluorescing, fade-retarding mounting medium. p0210 Another fluorescent tracer, Fluoro-Gold (FG), introduced by Schmued and Fallon in 1986, is also widely employed. FG can be used either alone or in combination with other fluorescent tracers, and it can be applied in combination with immunofluorescence detection of additional markers. The dye accumulates in small punctate structures, presumably lysosomes, in the cytoplasmic compartment of neuronal cell bodies. At higher concentration it can be seen to fill neurons completely with intense white/yellow fluorescence under UV illumination with a mercury lamp. This fluorescence is extremely resistant to bleaching, and in a living animal it may resist metabolic breakdown up to a year post injection. Its labeling intensity has made FG the gold standard for fluorescent labeling in rodents, particularly for multiple labeling in combination with other tracers. Due to its wide emission spectrum FG is less ideal in combination with other fluorescent tracers for retrograde double labeling when the purpose is to demonstrate collateralization, since it tends to mask the presence of the other fluorescent tracer in the same cells (Schmued and Fallon, 1986), or for application in confocal laser scanning instruments. However, the availability of an antibody against FG (Chang et al., 1990) makes it possible to take material studied previously in the fluorescence microscope to the electron microscope (Deller et al., 2000). FG is marketed by Fluorochrome, Inc. (Denver, CO) and its application is via mechanical injection or iontophoresis. The tracer is usually dissolved in cacodylate buffer, pH 7.3 (Schmued et al., 1990). Survival periods range from 1 week up to 1 year, and tissue fixation and sectioning can be conducted along standard neuroscience laboratory procedures. With long survival periods (more than 1 week), the retrogradely labeled neurons stand out with high contrast to the background. s0090 17.3.3.4 Dextrans p0215 Glover and coworkers published in 1986 a paper in which they described neuroanatomical tract tracing with a new tracer: dextran amine conjugated with the fluorescent dye rhodamine. This application of dextrans spawned an entire family of dextran-based tracers that have been successfully employed in the nonhuman primate model, notably dextran amine-Lucifer Yellow (LYD) (Chang, 1991), dextran aminetetramethylrhodamine (FR) (also called “Fluoro-Ruby”) (Schmued et al., 1990), fluorescein dextran (FD) (Nance and Burns, 1990), dextran-conjugated Alexa-488 (D-A488) (Rosenzweig et al., 2010) and, most importantly, biotinylated dextran amine (BDA) (Veenman et al., 1992). Contrary to the bidirectional transport of HRP, WGA,

and CTB, or the predominant retrograde transport of many other tracers (see sections above), dextrans are taken up and transported nearly exclusively in the anterograde direction. Unlike many other tracers, the mechanism of uptake is unknown (Reiner et al., 2000). However, it was found that BDA was rapidly taken up by somas within the injection site and transported anterogradely where it elegantly labeled the entire neuron, including the axon fiber, axon collaterals, terminal axon branches, and terminal boutons (Figure 17.12). The exquisite morphological detail of the dextran tracer method allows for individual axon reconstruction analysis (Ding et al., 2000; Borra and Rockland, 2011), detailed quantification of axon terminal projection fields (Lacroix et al., 2004; McNeal et al., 2010; Yoshino-Saito et al., 2010) as well as tracking terminal neuroplastic responses which accompany motor recovery following CNS injury including motor cortex resection (McNeal et al., 2010) or spinal cord transection (Rosenzweig et al., 2010). This characteristic has also made this tracer highly efficient, not only for single-tracing purposes, but also for combined applications since the detection of biotin is completely independent from, yet completely compatible with, immunocytochemistry. It was also found that BDA could be applied with few adaptations in electron microscopic neuroanatomical tracing (Wouterlood and JorritsmaByham, 1993) since the detection of transported BDA does not suffer from problems like penetration of antibodies into sections to the degree encountered with, for example, lectins. BDA is also well suited for doublelabel electron microscopic purposes. This wide versatility, together with the ease of application and the reliability of the tracer, has made BDA a popular neuroanatomical tracer. Biotinylated dextran amine conjugated to lysine (MW p0220 10 kDa, Invitrogen-Molecular Probes, Eugene, OR, USA) is dissolved as a 5% (rodents) or 10% (primates) solution in 10 mM phosphate buffer, pH 7.25 and injected under stereotaxic guidance into the brain. Both iontophoretic and mechanical injections have been reported (Reiner et al., 2000). Survival time is in proportion to the length of the projection under study. In the rat, BDA remains stable up to 4 weeks, but in the non-human primate it can remain stable from 3 to 6 months post injection (Lacroix et al., 2004; Yoshino-Saito et al., 2010). The tracer is compatible with a wide range of fixatives which implies that the choice for a particular fixative in multilabel experiments depends on the most demanding marker or additional immunocytochemistry. After fixation, the brain can be cut with any of the available sectioning methods. Sections with a thickness up to 400 mm can be processed. Thin sections (less than 40 mm) may be required when a second or third marker is visualized in addition to BDA to reduce background and improve the microscopic clarity of the reaction


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f0065 FIGURE 17.12

Example of a biotinylated dextran amine (BDA) injection site in the physiologically defined arm representation of M1 (a) in the rhesus monkey and resultant labeled fibers in the internal capsule (b). The inset in (b) shows a higher magnification of a group of labeled fibers in the internal capsule. (c) BDA-labeled fibers in the pyramidal decussation (PD) coursing into the lateral corticospinal tract (LCST) (see arrows) at the medullary–C1 junction. (d) A field of BDA-labeled axons and axon terminals in the spinal cord gray matter (Rexed’s laminae VII and IX) along with intricately defined terminal boutons (arrowheads). The asterisks indicate the location of motor neurons in lamina IX. The small inset in (d) shows in high magnification labeled boutons or swellings that indicate putative synaptic contacts. cs, central sulcus; gp, globus pallidus; ic, internal capsule; rtn, reticular thalamic nucleus. Scale bars: (a) 2 mm, (b) 1 mm, (c) 200 mm, (d) 50 mm.

products. Visualization of transported BDA is simple: reaction with a streptavidin–biotin complex (e.g. Vectastain kit, Vector, Burlingame, CA, USA) followed by incubation in a standard DAB–H2O2 mixture (Wouterlood and Jorritsma-Byham, 1993; Wouterlood et al., 1997). An alternative method of visualization is via a reaction of the transported BDA with streptavidin conjugated to a fluorochrome. Preparations obtained in this way can be studied in the fluorescence microscope or in a confocal instrument. s0095 17.3.3.5 Phaseolus vulgaris Leucoagglutinin (PHA-L) p0225 From the common kidney bean, Phaseolus vulgaris, several lectin subunits can be extracted which have either leucocyte agglutinating (L-subunit) or erythrocyte agglutinating (E-subunit) properties. A combination of four L-subunits produces a stable lectin, which, after deposition into the CNS, appears to be taken up nearly

exclusively by neuronal cell bodies and transported in the anterograde direction (Gerfen and Sawchenko, 1984). Uptake is thought to be receptor-mediated. Detection is via an antibody raised in rabbit or goat against the lectin, and this detection results in the visualization of the most exquisite details of the entire neuron, most importantly the fibers and axon collaterals, the terminal branches of the axon down to and including the terminal boutons (Figure 17.13). The fact that detection depends on an immunocytochemical procedure opened the way for combination of this tracing method with parallel immunocytochemical procedures (multiple staining) (Gerfen and Sawchenko, 1985) and with other tracing methods, for instance retrograde transport of a fluorescent tracer or even anterograde tracing with a noncompetitive tracer substance like BDA. The good compatibility of the two anterograde tracers, PHA-L and BDA, is extremely important because it opens up questions like convergence or divergence of projections


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17. CLASSIC AND CONTEMPORARY NEURAL TRACT-TRACING TECHNIQUES PHA-L – terminal labeling in dentate gyrus

PHA-L – overview

(a) BDA (fibers)

(b) parvalbumin (cells)

3D reconstruction

Contact

Green: BDA-fiber Red: parvalbumin dendrite

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f0070 FIGURE 17.13 (a) Low-magnification micrograph of a Phaseolus vulgaris leucoagglutinin (PHA-L) anterograde tracing experiment in rat

brain. Injection site, medial entorhinal cortex (m-EC), and labeling of terminal plexus (arrowheads) in the hippocampal region: subiculum (Sub), CA1, CA3 and molecular layer of dentate gyrus (DG); fh, hippocampal fissure. (b) High-magnification micrograph of PHA-L terminal labeling in the dentate gyrus, showing fibers and their characteristic swellings or boutons which are the presynaptic terminal elements of the neurons involved. (c) Two-channel fluorescence confocal image at medium magnification of the subiculum in a similar experiment; BDA used as anterograde tracer. Terminal BDA fiber labeling (488 nm channel, color-coded green). The neurons in the terminal zone are GABAergic interneurons, stained with an antibody against parvalbumin (568 nm channel, color-coded red). (d) 3D reconstruction after high-magnification confocal image acquisition of a BDA-labeled fiber in the subiculum with its terminal bouton (green), in close apposition with the dendrite of a parvalbumin-immunoreactive neuron (red).

for detailed study in one and the same experimental animal (Lanciego and Wouterlood, 1994; Lanciego et al., 1998). Combination of PHA-L or BDA as an anterograde tracer with a retrograde tracer opens ways to study in detail the anatomical connectivity of chains of neurons (Lanciego and Wouterlood, 2006). Contrary to its efficacy in rodents regardless of transport distances, anterograde labeling with PHA-L is primarily valuable for the study of short terminal projections in non-human primates. s0100 17.3.3.6 Combined Neuroanatomical Tract-Tracing Methods p0230 The inherent complexity of brain circuitry requires the design of sophisticated experimental paradigms enabling the visualization of multiple circuits in single sections. This particularly holds true when thinking in

terms of the complex networks of connections that typically characterize the brain of non-human primates. At present, up to three tracers can be used simultaneously in a single tissue section (Lanciego and Wouterlood, 2006) with as much as five tracers combined in a given experimental brain (Morecraft et al., 2001, 2002, 2007a). Moreover, the recent demonstration that dextran-conjugated Alexa-488 (D-A488) can successfully label corticospinal projection terminals following its injection into motor cortex in the nonhuman primate (Rosenzweig et al., 2010) suggests that it should be now possible to use up to six tracers in a single experiment. These tracers would include BDA, D-A488, FD, FR, LYD, and PHA-L. Strategies of this kind have a clear added value, since on the one hand an impressive amount of data dealing with how multiple circuits interact with each other is generated, and


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on the other hand the number of required experimental animals is markedly reduced. p0235 In the process of designing multitracer paradigms in non-human primates, two issues play a pivotal role in the final selection of tracers. First, a choice should be made depending on which tracers are best suited for their combined use within the experimental strategy. Second, tracer selection depends on which kind of visualization method is going to be applied (e.g. whether multiple colorimetric or multiple fluorescence detection methods are planned, or a combination of these). p0240 Regarding tracer selection, several points should be taken into consideration. First, the tracers should be fully compatible in terms of equivalent survival times so that all the selected tracers could be injected in one single surgical session avoiding repeated surgical procedures. Second, another desirable feature is that all the selected tracers allow for both the immunocytochemical and the immunofluorescence detection of the structures displaying labeling. Third, the availability of commercial antisera for tracer detectiondwhen requireddis another important demand. Antisera should be prepared in different animal species in order to prevent undesired cross-reactivity phenomena. Finally, with respect to stereotaxic placement in non-human primates, tracers should be selected that can be be pressure-injected using Hamilton syringes. In this regard, it should be borne in mind that brain coordinates required for stereotaxic surgery in primates are usually calculated by relying on ventriculography (in the same way as human neurosurgery before the arrival of MRI) and therefore the injection needle should be opaque under the Rx plate. Injection procedures other than pressure delivery (for example, microiontophoretic injections typically used for tracer delivery in rodents) are based on the use of glass micropipettes that cannot be properly visualized under Rx illumination, therefore increasing the risk of mistargeting. Although micro-iontophoretic application of the tracer is a useful choice when approaching small brain targets, the opposite is required for primates, in which selected target areas are often bigger. p0245 Keeping in mind these considerations, when attempting to trace up to three different brain circuits in one single tissue section of non-human primate material, a powerful combination of tracers is represented by the simultaneous use of BDA, FG, FB, DY, and CTB. The anterograde tracer BDA is a reliable tracer characterized by a broad range of survival times as previously mentioned. Indeed, BDA detection is a simple procedure enabling both the colorimetric form and the fluorescence form of visualization, with the latter taking advantage of the availability of a broad range of HRPor fluorochrome-labeled streptavidin conjugates. Furthermore, the visualization of BDA within a protocol that is not dependent on antisera facilitates the

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combination of BDA with the immunohistochemical detection of a host of other markers. Finally, a large number of vehicles (distilled water, phosphate buffer, and cacodylate buffer) can be used to deliver BDA mechanically through a Hamilton syringe. As mentioned, it is worth noting that the anterograde tracer PHA-L only transports adequately when delivered via microiontophoresis and therefore poses a serious limitation for its application in primates. Nevertheless, PHA-L can still be considered as an attractive second choice if the injection is to be located in the primate cerebral cortex. For retrograde tracing purposes, FB, DY, FG, and CTB have a wide range of survival times, can be pressure delivered, and detected either by immunocytochemistry or immunofluorescence. Indeed, FB, DY, and FG are inherently self-fluorescent under UV illumination, whereas several combinations of fluorochrome conjugates of CTB have recently been made available from Invitrogen-Molecular Probes (Leiden, The Netherlands). EXAMPLE OF A MULTIPLE TRACT-TRACING PARADIGM

s0105

In order to demonstrate the power and versatility of a p0250 multitracer experiment in non-human primates, we have chosen the combination of BDA þ FG þ CTB for illustrative purposes. Our experimental goal was to analyze whether pallidal afferents arising from the internal segment of the globus pallidus (GPi) that reach the substantia nigra pars compacta (SNc) innervate SNc neurons projecting either to the caudate nucleus or to the putamen of the primate Macaca fascicularis. Pallidonigral afferents were labeled after the deposit of the anterograde tracer BDA in GPi, whereas the delivery of the retrograde tracers FG and CTB into the putamen and caudate nucleus, respectively, identified the two different types of nigrostriatal-projecting neurons (a summary of the experimental plan is illustrated in Figure 17.14). All tracers were pressure-injected in a single surgical session. After a survival time of 2 weeks, the animals were perfused transcardially and the brains were removed and cryoprotected. Frozen sections (40 mm thick) were obtained with a sliding microtome and further divided in several series of adjacent tissue sections. Different series of sections were used for either multiple colorimetric detection (Figure 17.15) or for multiple fluorescence visualization (Figure 17.16). Multiple colorimetric detection relies on the use of three different peroxidase substrates, first comprising the BDA visualization using a nickel-enhanced diaminobenzidine chromogen (BDA-labeled axons stained blue– black), followed by a regular diaminobenzidine solution for the visualization of CTB-containing neurons in brown color. Finally, FG-labeled neurons were stained purple by means of the peroxidase substrate Vector-


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CTB injection site Retrograde transport

Caudate nucleus

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Nigrostriatal pathway Putamen

GPe

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Anterograde transport

SNc

Pallido-nigral pathway

SNr

f0075 FIGURE 17.14

Summary of the experimental plan conducted in primates comprising the injection of three tracers into three different basal ganglia nuclei. The goal was to identify projection neurons within the substantia nigra pars compacta (SNc) innervating the caudate nucleus and the putamen, those projection neurons in turn receiving afferents arising from the internal division of the globus pallidus (GPi). For this purpose, the retrograde tracers Fluoro-Gold (FG) and cholera toxin b subunit (CTB) were delivered within the putamen and the caudate nucleus, respectively. Next, the anterograde tracer biotinylated dextran amine (BDA) was injected in GPi. After 2 weeks of survival time, the primates were perfused and the structures displaying labeling were identified by means of either multiple immunoperoxidase or with multiple immunofluorescence protocols.

VIP. Multiple colorimetric detection resulted in permanent microscopical preparations that were stable in time. This approach is particularly well suited for the mapping of all structures displaying labeling (Figure 17.15). Overlapping areas between pallidonigral afferents and different subtypes of nigrostriatalprojecting neurons could easily be identified and plotted in camera-lucida diagrams, therefore supporting the elucidation of the overall pattern of relationships between identified structures. By contrast, quite different images are obtained when conducting multiple fluorescence visualization. The inspection of the sections under the confocal microscope at high resolution has a clear added value since it allows the identification of presumptive contacts between pre- and postsynaptic structures (Figure 17.16). Although the unequivocal identification of true synapses between axon terminals

of pallidothalamic projections and cell bodies of nigrostriatal-projecting neurons cannot be accomplished unless performing demanding triple pre-embedding ultrastructural examination (Reiner and Anderson, 1993), our view is that high-resolution confocal laser scanning is a simple procedure that properly fills the gap between light and electron microscopy, at least to some extent (Wouterlood et al., 2003, 2007, 2008b). Finally, it is worth noting that multiple immunofluo- p0255 rescence detection exhibits a clear added value over multiple colorimetric detection when trying to ascertain the chemical fingerprint of projection neurons, the latter identified by means of retrograde tracing. In other words, one might be interested in elucidating which kind of neurotransmitter is used by a given population of projection neurons. This is exemplified in Figure 17.16, in which the relationship between pallido-nigral fibers


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f0080 FIGURE 17.15

Detection of three transported tracers in one single histological section by means of three different peroxidase substrates. Coronal section through the primate substantia nigra showing pallido-nigral fibers and terminals, anterogradely labeled with BDA and stained in black color with the peroxidase substrate nickel-enhanced diaminobenzidine (DAB-Ni). Next, nigral neurons innervating the caudate nucleus were retrogradely labeled with CTB and visualized in brown color using a regular solution of diaminobenzidine (DAB). Finally, neurons projecting to the putamen were retrogradely labeled with Fluoro-Gold and stained in purple color, taking advantage of the peroxidase chromogen VectorÒ VIP. At low magnification (a, b), several areas of overlap between anterogradely labeled pallido-nigral axons and different subtypes of nigrostriatal neurons were clearly identified. At higher magnification (c), presumptive contacts between afferent fibers and efferent neurons were noticed. Scale bars: (a) 1000 mm, (b) 250 mm, (c) 80 mm.

reaching the substantia nigra (BDA-labeled) and nigrostriatal neurons projecting to the putamen (labeled with FG) is further implemented by demonstrating that nigrostriatal-projecting neurons have a dopaminergic nature. For this purpose, FG-labeled neurons are detected using a primary antibody against FG followed by an Alexa FluorÔ 488-conjugated secondary antibody (green channel). Next, dopaminergic neurons are indirectly detected using a primary antibody against tyrosine hydroxylase and then with an Alexa FluorÔ 546-conjugated secondary antibody (red channel). Finally, BDA-positive fibers are visualized by incubating the sections with Alexa FluorÔ 633-conjugated streptavidin (infrared channel). s0110 EXAMPLE OF CO-EXPRESSION OF NEUROTRANSMITTER AND CELL MARKERS IN INDIVIDUAL AXONS, AXON TERMINALS, AND CELL BODIES

p0260

Fluorescent compounds like FG usually produce only retrograde labeling of neuronal cell bodies, with the exception of FB that also produces anterograde and retrograde labeling of axons (see Section 17.3.3.3, above). Yet, by using the anterogradely transported BDA in a combined experiment, and capitalizing on the option to react the transported BDA with fluorescence-tagged streptavidin, one can study the innervation of

retrogradely labeled neurons by anterogradely labeled fibers. One step further is to use retrograde fluorescence tracing to identify the cell body of neurons belonging to a particular projection, inject intracellularly a fluorescent dye into the retrogradely labeled cell body, and follow up with analysis of the distribution of contacts between fibers labeled with the anterograde tracer over the dendrites of the intracellularly filled cell. Such experiments were first performed in 1991 and 1992 (Wouterlood, 1991; Wouterlood et al., 1992) whereas double anterograde neuroanatomical tracing combined with retrograde transport was further refined by Lanciego et al. (1998). These fluorescence methods were developed in the p0265 1980s and 1990s in parallel with technological advances leading to confocal laser scanning microscopy and related computer image processing. As a consequence of the digital image acquisition and image revolution, the fluorescence methods in combination with digital image acquisition procedures, followed up with computer imaging, have taken a dominant position in neuroanatomical tract tracing. We provide here a stateof-the-art example of multifluorescence work done in collaboration between various European and American laboratories (Figure 17.17). The scientific question was to distinguish in medial temporal cortex of the rat between extrinsic and intrinsic glutamatergic innervation


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f0085 FIGURE 17.16

The use of confocal microscopy to ascertain the chemical phenotype of a given type of projection neuron (identified by retrograde transport of Fluoro-Gold) receiving pallidal innervation (axons and terminals anterogradely labeled with BDA). The “blue” channel shows pallidal axons and terminals spreading over the substantia nigra pars compacta. These BDA-labeled fibers were visualized using streptavidin coupled with Alexa FluorÔ 546. The “red” channel shows neurons positive for tyrosine hydroxylase (TH; an indirect marker for dopaminergic neurons). Neurons positive for TH were visualized with a primary antibody against TH raised in mouse followed by a Alexa FluorÔ 633 goat anti-mouse secondary antibody. Finally, the “green” channel illustrates neurons retrogradely labeled with Fluoro-Gold. Neurons positive for Fluoro-Gold were identified with a rabbit anti-FG primary antibody, followed by incubation with a goat anti-rabbit bridge antibody coupled with Alexa FluorÔ 488. Only a subpopulation of TH1 neurons displayed immunofluorescence for Fluoro-Gold. Scale bar (a–d) 120 mm. At higher magnification (e–h) presumptive contacts between pallido-nigral terminals and nigro-putaminal projecting neurons showing a dopaminergic phenotype can be elucidated (arrowheads). Scale bar (e–h) 40 mm.

further characterized by the presence of the calciumbinding protein calretinin (Wouterlood et al., 2008a). The extrinsic fibers were labeled via anterograde neuroanatomical tracing with BDA. The BDA was visualized using the fluorescent tag Alexa FluorÔ 594 conjugated to streptavidin. Identification of these fibers as glutamatergic was accomplished via an antibody raised in rabbit against vesicular glutamate transporter 2 (VGLUT2), a synaptic vesicle-bound protein that accumulates glutamate across the vesicle’s membrane into the synaptic vesicle. The secondary antibody here was goat–anti-rabbit conjugated with the fluorescent tag Alexa FluorÔ 488. Co-identification of these fibers as containing calretinin was achieved via a mouse–anti-calretinin antibody detected with goat–anti-mouse IgG conjugated with Alexa FluorÔ 633. Thus, a triple-label preparation had been made in the same section with the three different fluorescence labels corresponding with laser illumination wavelengths typical for confocal laser scanning: 488–594–633. With proper calibration, sequential mode scanning, signal filtering, and via post-acquisition image processing, the confocal scanning instrument is capable of determining co-expression of markers in individual

fibers and axon terminals. With this instrument, employing the three laser illumination wavelengths corresponding with the three fluorochromes, and after image processing and 3D reconstruction, proof of colocalization was established for two immunocytochemical markers in terminals belonging to tracer-labeled fibers (Figure 17.17) (Wouterlood et al., 2007, 2008b). Other rats were examined through a combination of retrograde tracing with FG (injection in medial temporal cortex) and immunofluorescence with calretinin antibodies to determine whether the cells of origin contained both the tracer and the specific marker. Thalamic neurons located in one of the midline nuclei (nucleus reuniens thalami) indeed contained retrogradely transported FG while simultaneously they immunostained positive with the anti-calretinin antibodies. However, with these tracing approaches we still could not distinguish interneurons with short axons that distribute within the medial temporal cortex. Interneurons notoriously tend to escape tracing with neuroanatomical tracing methods which, because of their manner of application, are best suited to study long, extrinsic projections. The interneuron issue was solved


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17.3. CONTEMPORARY APPLICATION OF EXPERIMENTAL TRACT TRACING IN NON-HUMAN PRIMATES 594 nm: BDA

488 nm: VGLUT2

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(c) 3D reconstruction

Overlay: 488-594-633 nm channels

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Green: VGLUT2 Red: BDA Blue: CR

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f0090 FIGURE 17.17

Triple fluorescence experiment in the rat: the anterograde tracer BDA injected in the nucleus reuniens of the midline thalamus and two additional markers: vesicular glutamate transporter 2 (VGLUT2) that labels synaptic vesicles in glutamatergic boutons and the calciumbinding protein calretinin (CR). Confocal images acquired in parallel channels at very high magnification of one spot in the medial temporal cortex. (a) The 488 nm laser channel (color-coded green) shows VGLUT2 immunoreactive boutons. (b) The 594 nm laser channel (color-coded red) shows a BDA-labeled fiber with several varicosities (boutons). (c) The 633 nm laser channel shows CR immunoreactive fibers and boutons. (d) Overlay image of all three channels. (e) 3D reconstruction of the triple-labeled bouton indicated with arrow in a, b, and c.

by a combination of fluorescence in situ hybridization (the probe was a messenger RNA that coded for VGLUT2) and immunofluorescence to identify calretinin-containing cell bodies. As we found doublepositive cells in the medial temporal cortex, our final conclusion was that both thalamic glutamatergic projection neurons and medial temporal cortical glutamatergic interneurons contribute to the innervation of this regional cortical area within a network of calretinincontaining neurons (Wouterlood et al., 2008a). s0115 EXAMPLE OF MULTIPLE ANTEROGRADE TRACER COMBINATIONS TO STUDY THE SPATIAL RELATIONSHIPS BETWEEN EFFERENT PATHWAYS

p0270

As described earlier in this chapter, the quest for tracing fiber pathways spans centuries of experimental investigation, going as far back as the early attempts using gross dissection, to the relatively crude microscopic degenerating techniques developed by Marchi (Marchi and Algeri, 1885). The development of tritiated amino acids unquestionably continued to add to our ability to

study the complex trajectories of efferent fiber pathways at the cortical as well as subcortical levels. However, the advent of contemporary tracers, particularly the dextrans, has led to our ability to trace numerous pathways in the same experimental brain while simultaneously determining the spatial relationships between these efferent pathways, including the degree of segregation or overlap shared among the studied pathways (Figure 17.18). This can be achieved by injecting different tract-tracing compounds into different regions of the same brain and then processing the tracers in adjacent tissue sections using immunohistochemistry. In the example illustrated, five different tracers were injected into five different cortical arm representations in the brain of a single rhesus monkey (Figure 17.19) (Morecraft et al., 2002, 2007a). The arm areas were located within the primary (M1), dorsolateral pre(LPMCd), supplementary (M2), rostral cingulate (M3), and caudal cingulate (M4) motor cortices. The tracer injections included PHA-L, fluorescein dextran (FD), Fluoro-Ruby (FR), BDA, and Lucifer Yellow–dextran


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f0095 FIGURE 17.18

Digital photomicrographs illustrating examples of immunohistochemically labeled fiber bundles in the midbrain crus cerebri in the rhesus monkey. In each panel the inset(s) show an enlarged image of the labeled fibers from the location depicted by the arrow (for experimental details see Morecraft et al., 2007b). (a) Representative section through superior levels of the midbrain showing BDA-labeled fibers (brown) and Lucifer Yellow–dextran (LYD)-labeled fibers (blue) following the injection of BDA into the primary motor cortex (M1) and LYD into the ventrolateral premotor cortex (LPMCv). Partial overlap characterized these pathways. (b) Representative section through superior levels of the midbrain showing BDA-labeled fibers (brown) and fluorescein dextran (FD)-labeled fibers (blue) following injection of BDA into LPMCv and FD into the LPMCd. Direct overlap characterized these pathways. (c) Representative section through mid-levels of the midbrain showing BDA-labeled fibers (brown) and FD-labeled fibers (blue) following injection of BDA into the LPMCd and FD into the supplementary motor cortex (M2). Direct overlap characterized these pathways. (d) Representative section through mid-levels of the midbrain showing BDA-labeled fibers (brown) and FD-labeled fibers following injection of BDA into the caudal cingulate motor cortex (M4) and FD into the rostral cingulate motor cortex (M3). Complete segregation characterized these pathways. Reproduced from Moorcraft et al. (2007a) with permission from John Wiley & Sons, Inc.

(LYD), respectively. Following a survival period to allow for tracer transport, the tissue was fixed, cryoprotected then frozen sectioned in the anterior/posterior commissural plane (i.e. horizontally). At every 500 mm interval, adjacent tissue sections were immunohistochemically processed for each tracer (Morecraft et al., 2007a). The data (tissue section outlines, gray and white matter borders, injection sites, and labeled fibers) in every processed tissue section were plotted using a microscope attached to a computer-assisted neuranatomical data collection station (Neurolucida, MBF Bioscience, Inc.)

and reconstructed to reveal the 3D relationships of the various descending pathways. At superior levels of the corona radiata p0275 (Figure 17.19a, b) fiber representation was found to be widespread with progressive overlap as the fiber bundles entered superior levels of the internal capsule (Figure 17.19e, f). Once within the internal capsule, the fiber bundles were located anterior to posterior. This widespread distribution has also been supported in recent clinical and human studies (e.g., Lang and Schieber, 2003; Fridman et al., 2004; Holodny et al., 2005; Newton et al., 2006; Raghavan et al., 2006;


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Schaechter et al., 2008; Sullivan et al., 2010). As the bundles coursed inferiorly, their overlap progressively increased and they were found as a group to gradually shift posteriorly. Given the potential of non-human primate motor corticofugal pathway homologies with the human brain (Morecraft et al., 2002), these findings provide strong support for the suggestion that the upper limb motor recovery is likely to decrease as a lesion site occupies, progressively, the cerebral cortex, corona radiata, superior internal capsule, and inferior internal capsule (Shelton and Reding, 2001; Schiemanck et al., 2008). Likewise, these data suggest that lesions located more posteriorly in the posterior limb of the internal capsule are likely to lead to more severe motor deficits than lesions located more anteriorly in the posterior limb. This hypothesis is being based on the fact that posterior injury would likely involve the corticofugal projection from M1 which gives rise to the most dense corticofugal projection, including the most robust cortico-spinal projection. It is noteworthy that this hypothesis received support from a recent clinical study examining the effects of small lesions seated in different regions of the posterior limb in human stroke patients (Wenzelburger et al., 2005). Progressing inferiorly into the midbrain, the fiber pathways shifted to a medial– lateral orientation (Figure 17.19i–k). In this location fibers from M1 were positioned centrally and fibers from the rostral cingulate motor cortex (M3) were positioned medially. The remaining fiber bundles (from M2, LPMCd, and M4) were located between. These findings support classical (Bucy and Keplinger, 1961) and recent clinical observations in human stroke patients with pure midbrain infarction (Kim and Kim, 2005) which suggest corticofugal fiber representation may be more widespread in the human peduncle than currently appreciated. For example, in the Kim and Kim study,

=

Representative serial sections of a fiber pathway f0100 experiment through the rhesus monkey corona radiata (a–d), internal capsule (e–h), and midbrain (i–n) following an injection of Phaseolus vulgaris leucoagglutinin (PHA-L) into the arm area of M1 (dark blue), an injection of fluorescein dextran (FD) into the arm area of dorsolateral premotor cortex (LPMCd) (green), and injection of Fluoro-Ruby (FR) into arm area of M2 (red), an injection of biotinylated dextran amine (BDA) into the arm area of M3 (light blue) and an injection of dextran amine-Lucifer Yellow (LYD) into the arm area of M4 (yellow) (from Morecraft et al., 2002, 2007a). Illustrated are the corresponding trajectories of the five pathways. Each descending fiber bundle is identified by a color-coded outline, and the cortical origin of each bundle is identified in the corresponding key. Note, posterior is oriented toward the bottom of panel I (sections a–h) and dorsal is oriented toward the top of panel II (sections i–n). In the cortical sections, the plane of cut is established through the anterior–posterior commissural plane (see section h on medial hemispheric wall–top right of panel). Reproduced from Moorcraft et al. (2002) with permission from Oxford University Press and from Moorcraft et al. (2007a) with permission from John Wiley & Sons Inc.

FIGURE 17.19

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38 of 40 patients followed in this study eventually attained “functional levels” of motor recovery (Morecraft et al., 2007a). Finally, although “locked-in syndrome” is more commonly associated with bilateral pontine infarction, these non-human primate findings suggest that the midbrain form of “locked-in syndrome” may be the consequence of bilateral injury to the most inferior region of the crus cerebri, since fiber organization in this location (i.e. the midbrain–pontine isthmus) is characterized by extensive overlap of all the descending motor pathways (Figure 17.19n).

s0120 17.3.4 Transneuronal Tracers p0280

A major landmark event in the evolution of tracttracing methods has been the development of transneuronal tracers (i.e. markers that are transferred specifically between connected neurons and allow for the visualization of entire functional neuronal networksdfirst-order, second-order, third-order neurons, etc.). The search for transneuronal transfer of substances was initiated as an attempt to clarify the mechanisms mediating trophic effects of neurons upon functionally connected cells. Such effects had been suggested by the phenomena of anterograde transneuronal degeneration of neurons following the death of their target cells and retrograde transneuronal degeneration occurring after deafferentation (Cowan, 1970). Two different types of traffic of substances were demonstrated: a specific one, involving transneuronal transfer between synaptically connected neurons and a nonspecific one, that is, local spread (transcellular transfer) consisting in the exchange of substances between neighboring, but not synaptically connected, glial cells and neurons (Grafstein and Forman, 1980). p0285 While both mechanisms represent a way for neurons to exchange information with their environment, only transneuronal transfer is specific and can be used to identify synaptically connected neurons. In other words, for a marker to be a reliable transneuronal tracer, transneuronal transfer should not be accompanied by local spread, which is a source of false-positive results. Therefore, in the long search for effective transneuronal tracers, validation of transneuronal markers has necessarily required a thorough evaluation of the specificity and extent of their transmission in experimental models of known connectivity. p0290 The development of these methodologies has involved first the use of some conventional tracers, and more recently the introduction of much more sensitive techniques, based on the use of viruses as transneuronal tracers. The purpose of this section is to provide a historical overview of the development of these new methodologies, focusing on viral transneuronal tracing techniques,

and to describe their different properties and examples of current applications in non-human primates. Among the viral transneuronal tracing methods p0295 currently available, the most recent one, based on the use of rabies virus, introduced by Ugolini in 1995, is also the most powerful one, since it makes it possible to trace neuronal connections across a practically unlimited number of synapses. Using this technique, polysynaptic neuronal networks that innervate a given CNS site or a single muscle, and which form the substrate of specific behaviors, can be mapped in their entirety, step by step, by studying the progression of the viral tracer at different time points after their injection (Ugolini, 2010, 2011). Viral transneuronal tracing studies, particularly in p0300 non-human primates, should be promoted, since the results that they provide form the “gold standards” for the design and interpretation of functional imaging investigations in humans. Notably, only viral transneuronal tracers can visualize functional neuronal networks at the highest resolution (at the neuronal level) (Figure 17.20) and in their entirety, regardless of their level of activation. Since the results obtained in nonhuman primates, particularly macaques, can be immediately translated to the human brain, the extremely refined knowledge of the organization of neuronal networks provided by viral transneuronal tracing studies in these species is of paramount importance for validating non-invasive imaging techniques and for improving the diagnosis and treatment of neurological diseases in humans. 17.3.4.1 Transneuronal Tracing Using Conventional Tracers

s0125

The first conventional marker used as transneuronal p0305 tracer was tritium-labeled proline, an amino acid, which is transferred exclusively in the anterograde direction. Its use enabled the classical demonstration of ocular dominance columns in the visual cortex in non-human primates (Wiesel et al., 1974). Since this tracer is membrane permeable but does not bind to neuronal receptors, its transfer largely involves passive diffusion from the transporting fibers to their target regions. In contrast, uptake and transneuronal transfer of the other two conventional markers which have been used as transneuronal tracers (i.e. WGA–HRP and tetanus toxin fragments) is receptor-mediated. Their main receptors are two types of gangliosides (the disialoganglioside GD1b and the trisialoganglioside GT1) for tetanus toxin fragments (Bizzini, 1979), and n-acetylglucosamine and sialic acid for WGA–HRP (Trojanowski, 1983). Detection of WGA–HRP is based on histochemical visualization of HRP, as reviewed earlier (Mesulam, 1982). For tetanus toxin fragments, detection of transneuronal labeling has been based on immunohistochemistry or coupling


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(a) Differences in transneuronal labeling obtained with conventional tracers (wheat germ agglutinin–horseradish peroxidase, WGA–HRP) and neurotropic viruses (i.e. rabies virus and alpha-herpesviruses (Herpes simplex virus type 1, HSV 1; Pseudorabies virus, PrV)). Modified from Kuypers and Ugolini 1990. With WGA–HRP (a), only a small amount of the tracer is transferred from first-order neurons (1 ) to second-order neurons (2 ), resulting in weak transneuronal labeling. Viruses function as self-amplifying markers (b, c): transfer to second-order neurons (b, 2 ) is followed by viral replication, producing intense transneuronal labeling (c, 20 ). (b–d) Examples of transneuronal labeling with rabies virus: thirdorder (3 ) corticofugal (layer V) neurons in cingulate cortex of nonhuman primates, labeled by retrograde transneuronal transfer of rabies virus from the lateral rectus muscle (G. Ugolini). Details from b are enlarged in c,d. Rabies transneuronal labeling is Golgi-like, visualizing the complete arborization of pyramidal neurons (b) (i.e. basal dendrites, dendritic spines (d) and even the distal end of apical dendrites ascending to the cortical surface (c)). Combination of rabies virus immunoperoxidase and Cresyl Violet counterstaining (see Ugolini et al., 2006). Scale bars: (b) 120 mm; (c, d) 40 mm. Reprinted from Kuypers and Ugolini (1990) with permission from Elsevier.

the fragments with strong isotopes or reporter genes (Buttner-Ennever et al., 1981; Horn and ButtnerEnnever, 1990; Perreault et al., 2006). p0310 Contrary to tritiated proline, tetanus toxin fragments and WGA–HRP are transported bidirectionally (Harrison

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et al., 1984). In the visual system, for example, WGA–HRP has been combined with tritiated proline for double anterograde transneuronal labeling, for a simultaneous demonstration of the ocular dominance columns related to the left and right eye (Horton and Hocking, 1996). With WGA–HRP, transneuronal labeling does not always occur; moreover, it may be intense only when secondorder neurons receive the tracers anterogradely or bidirectionally; a drawback is that bidirectional transfer hinders interpretations of the precise routes of labeling (Orioli and Strick, 1989). A common limitation of WGA–HRP and tetanus toxin fragments is the fact that retrograde transneuronal transfer is very inefficient: it occurs only if first-order neurons are filled with great quantities of the tracer. Even in these conditions, only small amounts of the tracer cross synapses from firstorder to second-order neurons (Figure 17.20a). As a result, retrograde transneuronal labeling with these conventional markers is very weak and can be detected only in some groups of second-order neurons, while thirdorder neurons cannot be visualized (Harrison et al., 1984; Porter et al., 1985; Kuypers and Ugolini, 1990). In primates, for example, retrograde transneuronal transfer of WGA–HRP from the lateral rectus muscle labeled only two second-order cell groups known to make inhibitory synaptic contacts with abducens motoneurons, whereas the other premotor cell groups were left unlabeled (Porter et al., 1985). With tetanus toxin fragments, transneuronal labeling may be more extensive, but it disappears in individual second-order premotor pathways depending on time (Horn and Buttner-Ennever, 1990). Another major drawback of tetanus toxin fragments is the fact that they inevitably spread from the injected muscle to many other muscles, such that retrograde transneuronal labeling cannot be confined to the innervation of a single motoneuron pool (Buttner-Ennever et al., 1981; Horn et al., 1995; Perreault et al., 2006). Because of these limitations, these classical retrograde transneuronal tracers are of limited value. 17.3.4.2 Viral Transneuronal Tracers

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The viruses used as transneuronal tracers are two p0315 alpha-herpesviruses (herpes simplex virus type 1: HSV 1, and pseudorabies: PrV) (Kuypers and Ugolini, 1990; Loewy, 1995; Ugolini, 1995b, 1996) and a rhabdovirus, that is, the fixed Challenge Virus Standard (CVS) strain of rabies virus (Tang et al., 1999; Ugolini, 1995a, 2010, 2011; Graf et al., 2002; Morcuende et al., 2002; Ugolini et al., 2006; Prevosto et al., 2009, 2010, 2011). Contrary to conventional tracers, they produce intense transneuronal labeling, because they replicate in recipient neurons (Figures 17.20, 17.21, and 17.22). Since they are fully competent viruses, their experimental use requires specific training and, in the case of rabies virus, prior vaccination. Moreover, experiments must be


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Differences in uptake of rabies virus by “slow” and “fast” motoneurons (MNs) (a,b, e–j) and example of retrograde transneuronal labeling of second-order neurons innervating fast MNs (c, d) at 2.5 days after injection of rabies virus into the distal (1) and central (2) part of the lateral rectus (LR) muscle (from Ugolini et al., 2006). (a, b) Selective uptake by slow MNs after distal injection (1) and by fast MNs after central injection (2). (b) Rabies injection area (red) in the LR muscle, visualized by rabies immunolabeling, showing that rabies virus does not spread within the muscle. Black dots: distribution of synaptophysin-positive terminals. (c,d) Examples of rabies retrograde transneuronal labeling of second-order neurons after central injection (2 in a, b) (uptake by fast MNs): transneuronal labeling in the contralateral magnocellular medial vestibular nucleus (MVmc, excitatory neurons of vestibulo-ocular reflex pathways to fast LR MNs, enlarged in d) and in the dorsal paragigantocellular reticular formation (DPGi, inhibitory burst neurons of saccade pathways). Rabies virus immunoperoxidase labeling is combined with Cresyl Violet counterstaining. (e, f) Three-dimensional reconstructions of the abducens (VI) nucleus, showing the differences in topography of retrogradely labeled slow MNs (e) and fast MNs (f) after injections in (1) and (2) (a, b). Dark blue outlines: VI nucleus and emerging roots of the VI nerve. Large dots: MNs cell bodies; small dots in e: labeled dendrites of slow MNs. Light blue outlines: descending limb of the facial nerve (VIIn). Green outlines: genu (g) and ascending limb of VIIn. Gray vertical lines (right): midline. Yellow outlines: brainstem dorsal surface. In contrast to fast MNs (f), slow MNs (e) are concentrated at the periphery of the VI nucleus and within the medial longitudinal fasciculus (MLF) (arrow in e). G–J) Examples of dual color immunofluorescence for rabies virus (FITC, green) and choline acetyltransferase (CAT), a motoneuron marker (Cy3, red) in the VI nucleus. Fast MNs in g,h; slow MNs in i,j. Rabies-infected MNs (g, i) remain viable, since they express CAT antigen (h, j) at normal levels (arrows). They are intermixed with unlabeled MNs (CAT-positive but rabies-negative). MVpc, medial vestibular nucleus, parvocellular; LV, lateral vestibular nucleus. Scale bars: (c) 600 mm; (d) 200 mm. (e, f) 400 mm; (g–j) 50 mm. Reproduced from Ugolini et al. (2006) with permission from John Wiley & Sons, Inc.

carried out at the appropriate biosafety containment level (2 or 3 depending on national regulations) (Ugolini, 1995a, 1995b, 1996; Enquist and Card, 1996; Kelly and Strick, 2000). s0135 ALPHA-HERPESVIRUSES (HERPES SIMPLEX VIRUS TYPE 1 AND PSEUDORABIES)

p0320

The first major step forward in the development of more sensitive and reliable transneuronal tracers was the introduction by Ugolini et al. (1987) of the retrograde

transneuronal tracing method based on the use of herpes simplex virus type 1 (HSV 1). HSV 1 is a DNA virus belonging to the alpha-herpesviruses family, which commonly causes cold sores in humans (Ugolini, 2010, 2011). By studying the propagation of HSV 1 from the hypoglossal nerve and limb nerves in rodents, these authors demonstrated that HSV 1 could serve to trace neuronal connections transneuronally across at least two synapses (Ugolini et al., 1987, 1989; Kuypers and Ugolini, 1990; Ugolini, 1992, 1995b, 1996, 2010). Because


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f0115 FIGURE 17.22 Retrograde transneuronal transfer of rabies virus after intracortical injections of a mixture of rabies virus and the conven-

tional tracer cholera toxin B fragment (CTB) in macaque monkeys (from Prevosto et al., 2010). (a) Summary of the pathways of rabies transneuronal transfer to the cerebellum after injection of the rabies–CTB mixture into the left ventral lateral intraparietal area (LIPv) or medial intraparietal area (MIP) of the intraparietal sulcus (IPS): 1 (black), first-order neurons (visualized by CTB) in the ipsilateral thalamus (white dots in e) and cortical areas. 2 (blue), second-order neurons labeled transneuronally (rabies virus) at 2.5 days in the contralateral cerebellar nuclei, in the ipsilateral thalamic nuclei and reticular thalamic nucleus, and in the contralateral thalamic nuclei (the latter reflecting projections to IPS areas of the right hemisphere). 3 (red), third-order neurons labeled at 3 days in the contralateral cerebellar cortex (Purkinje cells, PCs), and contralateral reticular thalamic nucleus. Anterograde transneuronal transfer (e.g. to the pontine nuclei) did not occur (X, violet). (b) Coronal sections showing the center of the injection area (red outlines), visualized by CTB immunoperoxidase at 2.5 days after injection of the rabies/CTB mixture into LIPv or MIP. (c, d) Photomicrographs of adjoining sections at the LIPv injection site, immunolabeled for CTB and rabies virus (same level as in b). Note that the injection area is easily identifiable with CTB (d) but not with rabies virus (c) at 2.5 days, because of strong rabies


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of its great sensitivity, transneuronal tracing with HSV 1 was rapidly adapted worldwide for the study of neuronal circuits, particularly in non-human primates (e.g. Zemanick et al., 1991; Hoover and Strick, 1993; Middleton and Strick, 1994). p0325 The other alpha-herpesvirus that is used for transneuronal tracing is PrV (also called herpes virus suis, since pigs are its natural host) (Loewy, 1995; Ugolini, 1995b, 1996, 2010). PrV is very similar to HSV 1 in structure and properties, and should not be confused with rabies virus (which is a negative-strand RNA virus belonging to the lyssaviruses family, and has completely different characteristics, Ugolini, 2010, 2011, see below). In rodents, both HSV 1 and PrV infect all categories of neurons (motoneurons, sensory neurons, and autonomic neurons) in the central and peripheral nervous systems. Because of their greater affinity for small sensory neurons and autonomic neurons than for motoneurons (Rotto-Percelay et al., 1992; Ugolini, 1992), PrV and HSV 1 have been found especially valuable for studying autonomic innervations of peripheral organs in rodents (Loewy, 1995). However, they are unable to infect primates after peripheral injections (Ugolini, 1995b, 2010). In contrast, HSV 1 propagates very efficiently by transneuronal transfer after injection directly into the CNS in primates (Zemanick et al., 1991; Hoover and Strick, 1993; Middleton and Strick, 1994; Ugolini, 1995b, 2010). Two HSV 1 strains containing specific mutations have been shown to exhibit interesting differences in the preferential direction of transfer following intracortical injections in primates: strain H129 is transferred mainly

in the anterograde direction, whereas strain McIntyreB is transferred almost exclusively in the retrograde direction (Zemanick et al., 1991; Hoover and Strick, 1993; Kelly and Strick, 2003; Ugolini, 2010). While the use of the McIntyre-B strain for retrograde transneuronal tracing has been recently discontinued in primates, due to the much greater specificity and efficiency of rabies virus, the H129 strain continues to be the only available viral tracer for anterograde transneuronal tracing, which complements the MRI-detectable manganese tracing (Saleem et al., 2002). Limitations of HSV 1 and PrV are the fact that they p0330 induce rapid neuronal degeneration and intense inflammatory response, and can propagate also via spurious (cell-to-cell) spread to glial cells and neurons (Ugolini et al., 1987; Loewy, 1995; Ugolini, 1995b, 1996). Moreover, when PrV is injected into the CNS, its uptake can also involve passing fibers crossing the injection site, although less prominently than some conventional tracers (Chen et al., 1999). Neuronal degeneration is unavoidable with HSV 1 and PrV because it is induced by specific viral genes that are required for neurovirulence and that shut off protein synthesis in the host cells (Ugolini, 1995b, 2010). Local spread is a source of falsepositive results when using these viruses to study neuronal connections (Loewy, 1995; Ugolini, 1995b, 1996, 2010). Attempts to dissociate local spread from transneuronal transfer by genetic engineering (i.e. by deleting specific viral glycoproteins mediating viral entry) have been unsuccessful. Local spread of HSV 1 and PrV is dependent on the virus strain and dose, and the time

=

immunolabeling of short-distance projections neurons in the IPS. In c, shading in the white matter ventral to LIPv is background staining of an area of fibrosis due to multiple recording traces. LIPd, dorsal LIP; VIP, ventral intraparietal area. (e) Example of rabies immunolabeling in the caudal thalamus (þ2.7 from the interaural axis) at 2.5 days after injection of the rabies–CTB mixture into MIP. Left side is ipsilateral. White dots: first-order neurons (CTB), here in lateralis posterior (LP), anterior pulvinar (APul) and medial dorsal (MD) nuclei. Brown: rabies retrograde transneuronal labeling. Labeling in the thalamus provides an internal control for the number of synapses crossed by the rabies tracer: at this time point (2.5 days), transfer involves second-order neurons (2 : ipsilateral reticular thalamic nucleus, Rt left, and contralateral thalamic nuclei) and not third-order neurons (3 : contralateral Rt, see summary diagram in a). Third-order labeling (e.g. in contralateral Rt) is obtained only at 3 days. IPul, inferior pulvinar; LG, lateral geniculate; MG, medial geniculate; NPC, nucleus of the posterior commissure; SG, suprageniculate. (f–i) Second-order (2 ) labeling in the cerebellar nuclei at 2.5 days (MIP) (see summary figure in a). (g) 3D reconstructions of the cerebellar nuclei, showing the second-order (2 ) populations that target MIP. Reconstructions were made using Neurolucida by stacking together serial 50 mm sections (200 mm spacing). The image shows a ventral view (90 rotation from the coronal plane) of the ensemble of the cerebellar nuclei, that was flipped on the y-axis to illustrate the contralateral (right) cerebellum on the right. Cerebellar nuclei and labeled neurons (dots) are colorcoded (blue: dentate, D; green: interpositus posterior, IP; orange-brown: interpositus anterior, IA; magenta: fastigial, F). Note the predominantly contralateral distribution of transneuronally labeled neurons, with major inputs to MIP from IP and D and minor contributions from IA and F. Photomicrographs in f, h, and i: examples of rabies-immunolabeled second-order neurons (2 ) in the cerebellar nuclei (MIP, 2.5 days, see g); note the intense labeling in ventrolateral IP and D (boxed area in h is enlarged in i). (j–l) Third-order (3 ) labeling of Purkinje cells (PCs) in the cerebellar cortex at 3 days (MIP) (see summary figure in a). (j) 3D reconstructions of the cerebellar cortex (posterior view), showing the modular organization of PCs that target trisynaptically MIP (3 days). Cerebellar divisions and labeled PCs in each division are color-coded. Most labeled PCs are found in PML (paramedian lobule) and Crus II posterior (IIp), dorsal paraflocculus (DPFl), paravermal anterior lobe (AL) and simplex. D, dorsal; M, medial; L, lateral; V, ventral. Photomicrographs in k and l: examples of rabies immunolabeled PCs in the cerebellar paramedian lobule (PML). Rabies virus immunoperoxidase labeling (in c, e, f, h, i, k, l) is combined with Cresyl Violet counterstaining. The results illustrate the power of the rabies transneuronal tracing technology in providing a reliable, time-dependent visualization of entire functional neuronal circuits (here, the cerebellar cortical and nuclear modules to MIP implicated in adaptive control of visual and proprioceptive guidance of reaching, arm/eye/head coordination, and prism adaptation, for details see Prevosto et al., 2010). Scale bars: (b) 4000 mm; (c–f) 2000 mm; (h) 1000 mm; (i) 100 mm; (i, j) 50 mm. Reproduced from Prevosto et al. (2010) with permission from Oxford Journals.


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post injection (Loewy, 1995; Ugolini, 1995b, 1996, 2010; Enquist and Card, 1996). Therefore, it can be minimized by manipulating these experimental parameters, but transneuronal transfer is also reduced (Loewy, 1995; Ugolini, 1995b). Typically, the conditions necessary to minimize local spread (use of attenuated strains which induce less cytopathological changes and injection of low doses and short time points) do not make it possible to trace further than second-order neurons (Loewy, 1995; Ugolini, 1995b, 1996, 2010). s0140 RABIES VIRUS

p0335

Continuing the search for a more reliable and efficient retrograde transneuronal tracer, Ugolini also explored the propagation of other viruses and introduced in 1995 the use of rabies virus (the “fixed” CVS-11 strain) for retrograde transneuronal tracing (Ugolini, 1995a, 2008, 2010, 2011). She demonstrated that, contrary to HSV 1 and PrV, rabies virus propagates exclusively between connected neurons via transneuronal transfer at chemical synapses without inducing neuronal degeneration or non-specific spread regardless of the dose and time post inoculation (Ugolini, 1995a, 2010, 2011; Tang et al., 1999; Graf et al., 2002; Grantyn et al., 2002; Morcuende et al., 2002; Moschovakis et al., 2004; Ugolini et al., 2006; Prevosto et al., 2009, 2010, 2011). Because of its unique properties, transneuronal tracing with rabies virus is the only viral tracing technique that is entirely specific and makes it possible to identify neuronal connections across an unlimited number of synapses. Another major advantage of rabies virus with respect to the other viral tracers is the fact that it is transferred transneuronally in all mammals including primates, after injections into muscles or directly into the CNS (Grantyn et al., 2002; Kelly and Strick, 2003; Moschovakis et al., 2004; Rathelot and Strick, 2006; Ugolini et al., 2006; Prevosto et al., 2009, 2010, 2011) (Figures 17.21 and 17.22). p0340 Rabies virus is a small, bullet-shaped, single-strand negative RNA virus; its genome (less than 12 kb) encodes only five proteins (Schnell et al., 2010; Ugolini, 2010, 2011). The single external glycoprotein (G) located on the viral envelope has a pivotal role for uptake and transneuronal transfer, as its gene deletion or point mutation abolishes transneuronal propagation of rabies virus (Finke and Conzelmann, 2005; Schnell et al., 2010; Ugolini, 2010, 2011). Transneuronal transfer is mediated by binding of the rabies virus glycoprotein to neuronal receptors; the most likely candidate is the neuronal cell adhesion molecule (NCAM) (Lafon, 2005; Ugolini, 2008, 2011). Rabies virus receptors are ubiquitously distributed within the CNS. In fact, in all models studied to date, retrograde transneuronal transfer of rabies virus has never failed to label known pathways mediated by classical chemical synapses (“wiring transmission,”

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Fuxe et al., 2007), regardless of their neurotransmitters, including both excitatory and inhibitory populations (Ugolini, 1995a, 2008, 2010, 2011; Tang et al., 1999; Graf et al., 2002; Grantyn et al., 2002; Morcuende et al., 2002; Ugolini et al., 2006; Prevosto et al., 2009, 2010, 2011). Only some “volume transmission” pathways (locus coeruleus, Fuxe et al., 2007) were less extensively labeled in some models (Ugolini 1995a, Tang et al., 1999), which may suggest that these special types of chemical synapses might be less conducive to rabies virus compared to alpha-herpesviruses or some conventional tracers (Ugolini, 1010). INTRACELLULAR CYCLE AND DIRECTION OF s0145 TRANSFER After uptake, rabies virus is transported p0345

retrogradely to the cell bodies of first-order neurons, where it replicates. Initially, rabies immunolabeling is restricted to the neuronal cell body; later, it extends to distal dendrites, but not to axons (Ugolini, 1995a, 2008, 2010, 2011). The finding that centrifugal intracellular transport of rabies virus targets exclusively dendrites explains why transneuronal transfer of rabies virus occurs only in the retrograde direction (i.e. from cell bodies and dendrites of first-order neurons to terminals of second-order neurons on their surface) (Ugolini, 2008, 2010). Remarkably, rabies virus propagates exclusively by retrograde transneuronal transfer even after injection directly into the CNS (e.g. intracortical injection) (Kelly and Strick, 2003; Prevosto et al., 2009, 2010, 2011; Ugolini, 2010) despite the fact that intracortical inoculations provide equal possibilities of uptake and axonal transport in the anterograde and retrograde directions (Figure 17.22). In fact, most conventional tracers are transported bidirectionally after such injections. The exclusively retrograde direction and high speed of axonal transport and transneuronal transfer of rabies virus can only be explained by active axonal transport mechanisms. These are mediated by the rabies virus glycoprotein (that has been shown to confer axonal transport properties to pseudotyped lentiviruses; Finke and Conzelmann, 2005; Ugolini, 2010, 2011) and by the strong interactions of the rabies virus P protein with the dynein light chain LC8, which is involved in intracellular retrograde axonal transport (Jacob et al., 2000; Ugolini, 2008; 2010, 2011).

IDENTIFICATION OF THE ORDER OF CONNEC- s0150 TIONS As for the other viral tracers, studying the ki- p0350

netics of transfer of the rabies tracer at different time points after the injections is of paramount importance for identifying the order of connections, since transneuronal transfer is strictly time-dependent. After an initial 2 days, required for infection of first-order neurons, higher order neurons (i.e. second-order, thirdorder, etc.) are labeled stepwise at regular intervals


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(12 hours or more, depending on the system and the viral load) (Ugolini, 1995a, 2008, 2010, 2011; Tang et al., 1999; Graf et al., 2002; Grantyn et al., 2002; Morcuende et al., 2002; Moschovakis et al., 2004; Ugolini et al., 2006; Prevosto et al., 2009, 2010, 2011) (Figure 17.22). The stepwise visualization of successive synaptic relays is due to the time required for each cycle of viral replication in recipient neurons. Notably, the distance does not influence the speed of visualization of different pathways of the same synaptic order (i.e. short pathways are labeled at the same time as very long ones) (Ugolini et al., 2006; Ugolini, 2010). The reason is that axonal transport of rabies virus occurs at very high rates, whereas viral replication in recipient neurons takes much longer; this “cancels out” the differences in tracer accumulation that can result from transport distances when using conventional (non-replicating) tracers (Ugolini, 2008, 2010, 2011). Visualization of neuronal networks across 6–7 synapses is obtained within less than a week. Experimental animals show a normal behavior during the entire study. This is in keeping with the long asymptomatic period of rabies, even in the case of very virulent natural isolates (“street” strains) which are not used for transneuronal tracing (Ugolini, 2010, 2011). s0155

EXCLUSIVE UPTAKE BY MOTOR ENDPLATES AFTER

p0355 INTRAMUSCULAR INJECTIONS After injection into muscles, rabies virus enters the nervous system exclusively via the motor route, that is, sensory and autonomic neurons which also innervate the muscle are not infected (Tang et al., 1999; Graf et al., 2002; Morcuende et al., 2002; Ugolini et al., 2006). Thus, in the peripheral nervous system, rabies receptors are present exclusively on motor endplates, contrary to their ubiquitous distribution within the CNS (Ugolini, 2008, 2010, 2011). The exclusive entry of rabies virus via the motor route is explained by the presence at the neuromuscular junction (but not at peripheral sensory and autonomic endings) of both the nicotinic acetylcholine receptor (nAChR, mainly at the postsynaptic location) and NCAM (at the presynaptic site), which both bind rabies virus. Because of its postsynaptic location, nAChR cannot directly mediate viral uptake, but may improve its probability by concentrating rabies virus particles in front of motor endplates, thereby facilitating entry which is mediated by NCAM (Lafon, 2005; Ugolini, 2011). Since motoneurons are the only gateway for retrograde transneuronal transfer of rabies virus to the CNS, this technique makes it finally possible to unravel with great specificity the organization of polysynaptic descending motor pathways involved in the control of single muscles in primates (Rathelot and Strick, 2006; Ugolini et al., 2006; Ugolini, 2010, 2011). Notably, this could not be achieved using HSV 1 and PrV even in

susceptible species (rodents), because their concurrent transfer in sensory, autonomic, and motor pathways makes it impossible to dissociate with certainly the contribution of transneuronal labeling to motor, versus sensory and autonomic innervation (Ugolini et al., 1989; Ugolini, 1992, 2010). EXAMPLES OF APPLICATIONS OF RABIES TRANS- s0160 NEURONAL TRACING The potential of retrograde p0360

transneuronal tracing with rabies virus for identifying functional neuronal networks at the highest resolution is exemplified by its application to studies of oculomotor pathways in primates. In macaque monkeys, Ugolini et al. (2006) have injected rabies virus into either the distal or central portion of an extraocular muscle (the lateral rectus), which contains the motor endplates of two different populations of abducens motoneurons, innervating slow and fast muscle fibers, respectively (Figure 17.21). They have found that rabies virus does not spread within the muscle, since it infected exclusively “slow” motoneurons after distal muscle injections, whereas “fast” motoneurons were labeled only after central muscle injections. Retrograde transneuronal transfer of rabies virus from these two motoneuron populations to the respective second-order neurons has revealed striking differences in their innervation (Ugolini et al., 2006). At longer time points of transfer, retrograde transneuronal labeling has made it possible to identify higher order neurons of brainstem and cortical oculomotor networks (Grantyn et al., 2002; Moschovakis et al., 2004). One of these studies (Moschovakis et al., 2004) has shown the correspondence in frontal lobe oculomotor areas between the distribution of activation during oculomotor tasks, visualized by 2-deoxyglucose functional imaging, and the topography of oculomotor corticofugal neurons that innervate abducens motoneurons di- and trisynaptically, labeled transneuronally by rabies virus. Compared with functional imaging, rabies transneuronal tracing has the tremendous added value of revealing the finest details of corticofugal oculomotor neurons and neuron networks and showing the exact number of synapses involved in the transmission of oculomotor commands to individual eye muscles. In addition to the great impact of rabies transneuronal tracing on the understanding of neuronal circuits, these neuroanatomical studies of the early stages of rabies virus propagation into the CNS are providing new insights into the pathogenesis of rabies disease (see Ugolini, 2008, 2011).

COMBINATION OF RABIES TRANSNEURONAL s0165 TRACING WITH DETECTION OF NEUROTRANSMITTERS, CELL MARKERS, AND CONVENTIONAL TRACERS Since p0365

this chapter is devoted mainly to tracing techniques used in non-human primates, we describe in this section


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only the available combinations of rabies transneuronal labeling with other techniques. For combinations of HSV 1 and PrV with other techniques, protocols are only available for rodents (Loewy, 1995; Ugolini, 1995b, 1996; Enquist and Card, 1996). p0370 Because of the amplification of the signal provided by viral replication, retrograde transneuronal labeling obtained with rabies virus is very intense, particularly using the most recent immunoperoxidase protocols of Ugolini et al. (2006), and it is routinely combined with Cresyl Violet counterstaining of the same section. Rabies transneuronal labeling is Golgi-like, revealing the finest dendritic arborization of labeled neurons, including dendritic spines (Figure 17.20). Even after long-term infection, the CVS-11 strain of rabies virus does not cause morphological changes suggestive of neuronal damage: infected neurons show normal size and Nissl staining patterns and do not undergo apoptosis (programmed cell death) (Ugolini, 1995a, 2010, 2011; Tang et al., 1999; Graf et al., 2002; Grantyn et al., 2002; Morcuende et al., 2002; Ugolini et al., 2006). In fact, there is now substantial evidence that rabies virus has developed a multilevel strategy to prevent neuronal impairment: its replication does not involve host shutoff mechanisms, it has the ability (virulent strains) to prevent apoptosis by keeping gene expression beyond threshold and by interfering with proapoptotic factors, and it also displays immunoevasive strategies (Schnell et al., 2010; Ugolini 2010, 2011). Thus, infected neurons remain metabolically viable, and can still transport other tracers and express their neurotransmitters (Tang et al., 1999; Graf et al., 2002; Grantyn et al., 2002; Morcuende et al., 2002; Miyachi et al., 2006; Ugolini et al., 2006; Prevosto et al., 2009, 2010, 2011; Ugolini, 2010, 2011). As a result, it is possible to combine rabies immunolabeling with neurotransmitter immunohistochemistry and other tracers. p0375 Protocols for combined visualization of rabies transneuronal labeling and neurotransmitter or cell markers are based on dual immunofluorescence procedures; published protocols describe the combined visualization of rabies virus and choline acetyltransferase (used as marker for motoneurons and autonomic preganglionic neurons) (Tang et al., 1999; Graf et al., 2002; Morcuende et al., 2002; Ugolini et al., 2006) (Figure 17.21g–j), oxytocin (Tang et al., 1999), calbindin, parvalbumin, pleiotrophin and the neuronal form of nitric oxide synthase (Miyachi et al., 2006; Salin et al., 2009). p0380 Rabies virus immunolabeling can also be combined with conventional tracers. Morcuende et al. (2002) have combined retrograde labeling of rubrospinal neurons using the fluorescent tracer Fluoro-Ruby (red) with immunofluorescent (FITC, green) detection of rabies transneuronal labeling of rubrobulbar neurons which innervate orbicularis oculi motoneurons. More recently,

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Prevosto et al. (2009, 2010, 2011) have shown that it is possible to inject a mixture of rabies virus and the conventional tracer, cholera toxin B fragment (CTB, end concentration 0.03%) without altering the uptake of either tracer (Figure 17.22). This is a major methodological improvement, since this tracer combination makes it possible to visualize in the same experiment the injection area and direct projections (first-order neurons) with CTB, in combination with transneuronal labeling of higher order neurons by means of rabies virus (Prevosto et al., 2009, 2010, 2011; Ugolini, 2010, 2011). The major advantages of this rabies–CTB combina- p0385 tion are summarized in Figure 17.22. In these experiments, the rabies–CTB mixture was injected intracortically, into the medial intraparietal area (MIP) or into the ventral portion of the lateral intraparietal area (LIPv) in macaque monkeys (see Prevosto et al., 2010). Brains were processed at different post-injection time points (2.5 or 3 days), when transneuronal transfer involved second-order and third-order neurons respectively; alternative series of sections were treated for CTB immunoperoxidase, or rabies immunoperoxidase combined with Cresyl Violet counterstaining. Photomicrographs of neighboring coronal sections at the center of the LIPv injection area (2.5 days, second-order time point) processed for CTB and rabies immunolabeling are shown in Figure 17.22c, d. As shown in Figure 17.22c, defining the precise extent of the injection area with rabies immunolabeling alone is difficult, because transneuronal labeling includes short-distance projection neurons in neighboring portions of the intraparietal sulcus; moreover, the rabies tracer does not accumulate at the injection site, does not induce tissue damage and does not label glial cells (note the razor-sharp end of the labeling at the border between gray and white matter) (Prevosto et al., 2010; Ugolini, 2010). Conversely, CTB immunolabeling allows for a precise definition of the extent of the injection site (Figure 17.22d). Since CTB is likely to diffuse more than rabies virus, a safety measure is to delimitate the maximum extent of the injection, as it might eventually overestimate but not underestimate the effective rabies injection area (Ugolini, 2010). Another important advantage is that the rabies–CTB mixture allows for the simultaneous identification of first-order neurons (CTB) and higher order neurons (rabies virus) (Prevosto et al., 2009, 2010, 2011) (Figure 17.22). Because CTB is deliberately used at low concentration (0.03%) in order not to interfere with rabies virus uptake, CTB immunolabeling is only retrograde, but easily detectable and visualizes all known direct projections (first-order) to the injected CNS target (e.g. thalamo-cortical neurons, Figure 17.22a, e). In the same experiment, rabies immunolabeling allows for a precise identification of the second-order (cerebellar nuclei at 2.5 days, Figure 17.22a, f–i) and third-order


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populations (cerebellar cortex Purkinje cells at 3 day, Figure 17.22a, j–l). Importantly, CTB (0.03%) does not alter uptake of the rabies tracer, because transneuronal transfer occurs at the same rate and with the same efficacy as after injection of rabies virus alone (for details, see Prevosto et al., 2009, 2010, 2011; Ugolini, 2010). This tracer combination strategy has a clear added value, since it makes it possible to identify unequivocally firstand higher order projections in the same experiment, and on the other hand it reduces the number of animals required to complete a study.

s0170 p0390

17.4 CONCLUSIONS

There is a rich and fascinating history on the development of methods employed to trace neural pathways which have led to an unprecedented armamentarium of scientific tools presently available in experimental neurology. In the present chapter we have chosen, on a selective basis, to emphasize monumental achievements that have led to the technical development of neural tract tracing, while discussing in detail the powerful methods that are currently being applied to investigate neural pathways in the non-human primate brain. The wide-ranging capabilities and deployment of multiple procedures in the same experimental brain that have been developed and refined over the past 30 years have all but abolished the many limitations once placed on addressing neuroanatomical questions in experimental animals. Indeed, there is common purpose in the theoretical foundation of both experimental tract tracing and the application of diffusion tensor imaging (DTI), stemming from the straightforward objective to unmask nature’s secrets of cell-to-cell communication in the CNS with the main goal to understand, treat, and cure disabling neurological disorders. There is no doubt that technical breakthroughs and new applications will persist in experimental neural tract tracing and these findings will continue to lead the way as advances are steadily made with the new and exciting non-invasive neuroimaging methods. p0395 In many regards, while DTI tractography is currently an electrifying field of discovery with unquestionably important and unique applications, including in vivo study of human whole-brain systems, it has fundamental limitations such as the inability to differentiate efferent and afferent pathways, identify collateral pathways, determine the precise degree of fiber pathway convergence (i.e. overlap) and divergence, and follow interconnected chains of pathways to highlight a few. Furthermore, a typically sized imaging voxel will contain thousands of axons, with potentially complex geometry. It is without question that in the years to come we will witness a methodological revolution in

the development of DTI. More sophisticated data acquisition schemes, enabling more complex modeling and tractography approaches, will allow for increasingly fine-grained pathways to be estimated. However, the fundamental limitations of tractography are insurmountable by technical advances. It is therefore important to stress that tractography does not remove the need for tract-tracing experiments, but rather provides a complementary approach. The ability to actively trace individual axons is unique to tracer studies, but these studies must be done in animal experiments. The ability to estimate the course of anatomical pathways in brains in living human beings is unique to tractography, but technical limitations mean that these estimates can suffer from significant false positives and false negatives. The results obtained from the application of advanced, high-resolution tract-tracing techniques in experimental animals, and in particular non-human primates, will therefore remain critical. A consistent and unsurprising conclusion from many DTI studies is that tractography in the human brain confirms what had previously been established in studies on non-human primates. This reiterates the relevance of existing and emerging careful anatomical studies of connectional anatomy in non-human animals to studies in human subjects in health and disease. While the revolution in both tract tracing and imaging continues, the best research will be done with plenty of cross-talk and collaboration between the two communities.

References Alipour, M., Chen, Y., Jurgens, U., 2002. Anterograde projections of the motorcortical tongue area in the saddle-back tamarin (Saguinus fuscicollis). Brain Behav. Evol. 60 (2), 101–116. Bassetti, C.L., Jagella, E.C., 2006. Joseph Jules Dejerine (1849–1917). J. Comp. Neurol. 253, 823–824. Bentivoglio, M., 1996. 1896–1996: the centennial of the axon. Brain Res. Bull. 41 (6), 319–325. Bentivoglio, M., Kuypers, H.G., Catsman-Berrevoets, C.E., Dann, O., 1979. Fuorescent retrograde neuronal labeling in rat by means of substances binding specifically to adenine-thymine rich DNA. Neurosci. Lett. 12 (2–3), 235–240. Bentivoglio, M., Kuypers, H.G., Catsman-Berrevoets, C.E., Loewe, H., Dann, O., 1980. Two new fluorescent retrograde neuronal tracers which are transported over long distances. Neurosci. Lett. 18 (1), 25–30. Bilgen, M., Peng, W., Al-Hafez, B., Dancause, N., He, Y.Y., Cheney, P.D., 2006. Electrical stimulation of cortex improves corticospinal tract tracing in rat spinal cord using manganeseenhanced MRI. J. Neurosci. Methods 156 (1–2), 17–22. Bizzini, B., 1979. Tetanus toxin. Microbiol. Rev. 43 (2), 224–240. Borra, E., Rockland, K.S., 2011. Projections to early visual areas v1 and v2 in the calcarine fissure from parietal association areas in the macaque. Front. Neuroanat. 5, 35. Brown, A., 2003. Axonal transport of membranous and nonmembranous cargoes: a unified perspective. J. Cell. Biol. 160 (6), 817–821.


10017-JOHANSEN-BERG-9780123964601


REFERENCES

Bryantseva, S.A., Hoerber, J.K., 2012. Bidirectional transport of organelles: unity and struggle of opposing motors. Cell. Biol. Int. 36 (1), 106. Bucy, P.C., Keplinger, J.E., 1961. Section of the cerebral peduncles. Preliminary report. Arch. Neurol. 5, 132–139. Buttner-Ennever, J.A., Grob, P., Akert, K., Bizzini, B., 1981. Transsynaptic retrograde labeling in the oculomotor system of the monkey with [125I]tetanus toxin BIIb fragment. Neurosci. Lett. 26 (3), 233–238. Chang, H.T., 1991. Anterograde transport of lucifer yellow-dextran conjugate. Brain Res. Bull. 26 (5), 813–816. Chang, H.T., Kuo, H., Whittaker, J.A., Cooper, N.G., 1990. Light and electron microscopic analysis of projection neurons retrogradely labeled with Fluoro-Gold: notes on the application of antibodies to Fluoro-Gold. J. Neurosci. Methods 35 (1), 31–37. Chen, S., Ming, X., Miselis, R., Aston-Jones, G., 1999. Characterization of transsynaptic tracing with central application of Pseudorabies virus. Brain Res. 838 (1–2), 171–183. Clark, E., O’Malley, C.D., 1996. The Human Brain and Spinal Cord: A Historical Study Illustrated by Writings from Antiquity to the Twentieth Century. San Francisco: Norman Publishing. Cowan, W.M., 1970. Anterograde and retrograde transneuronal degeneration in the central and peripheral nervous system. In: Nauta, W.J.H., Ebbesson, S.O.E. (Eds.), Contemporary Research Methods in Neuroanatomy. Springer, New York, pp. 217–251. Cowan, W.M., Gottlieb, D.I., Hendrickson, A.E., Price, J.L., Woolsey, T.A., 1972. The autoradiographic demonstration of axonal connections in the central nervous system. Brain Res. 37 (1), 21–51. Deller, T., Naumann, T., Frotscher, M., 2000. Retrograde and anterograde tracing combined with transmitter identification and electron microscopy. J. Neurosci. Methods 103 (1), 117–126. Ding, S.L., Van Hoesen, G., Rockland, K.S., 2000. Inferior parietal lobule projections to the presubiculum and neighboring ventromedial temporal cortical areas. J. Comp. Neurol. 425, 510–530. Droz, B., 1979. Elucidation of axonal transport by radioautography. J. Histochem. Cytochem. 27 (11), 1510–1511. Dum, R.P., Strick, P.L., 1996. Spinal cord terminations of the medial wall motor areas in macaque monkeys. J. Neurosci. 16 (20), 6513–6525. Duque, A., Zaborszky, L., 2006. Juxtacellular labeling of individual neurons in vivo: from electrophysiology to synaptology. In: Zaborszky, L., Wouterlood, F.G., Lanciego, J.L. (Eds.), Neuroanatomical Tract-Tracing 3: Molecules, Neurons, and Systems. Springer, New York, pp. 197–236. Edwards, S.B., Hendrickson, A., 1981. The autoradiographic tracing of axonal connections in the central nervous system. In: Heimer, L., RoBards, M. (Eds.), Neuroanatomical Tract-Tracing Methods. Plenum, New York, pp. 171–205. Enquist, L.W., Card, J.P., 1996. Pseudorabies virus: a tool for tracing neuronal connections. In: Lowenstein, P.R., Enquist, L.W. (Eds.), Protocols for Gene Transfer in Neuroscience: Towards Gene Therapy of Neurological Disorders. John Wiley and Sons, Chichester, pp. 333–358. Finger, S., 1994. Origins of Neuroscience. Oxford University Press, Oxford. Fink, R.P., Heimer, L., 1967. Two methods for selective silver impregnation of degenerating axons and their synaptic endings in the central nervous system. Brain Res. 4 (4), 369–374. Finke, S., Conzelmann, K.K., 2005. Replication strategies of rabies virus. Virus Res. 111 (2), 120–131. Foster, M., assisted by Sherrington, C.S., 1897. A Textbook of Physiology, Part 3: The Central Nervous System, seventh ed. Macmillian & Co, London. Fridman, E.A., Hanakawa, T., Chung, M., Hummel, F., Leiguarda, R.C., Cohen, L.G., 2004. Reorganization of the human ipsilesional premotor cortex after stroke. Brain 127, 747–758.

395

Friedland, D.R., Los, J.G., Ryugo, D.K., 2006. A modified Golgi staining protocol for use in the human brain stem and cerebellum. J. Neurosci. Methods 150 (1), 90–95. Fuxe, K., Dahlstro¨m, A., Hoïstad, M., Marcellino, D., Jansson, A., Rivera, A., Diaz-Cabiale, Z., Jacobsen, K., Tinner-Staines, B., Hagman, B., Leo, G., Staines, W., Guidolin, D., Kehr, J., Genedani, S., Belluardo, N., Agnati, L.F., 2007. From the GolgiCajal mapping to the transmitter-based characterization of the neuronal networks leading to two modes of brain communication: wiring and volume transmission. Brain Res. Rev. 55 (1), 17–54. Gerfen, C.R., Sawchenko, P.E., 1984. An anterograde neuroanatomical tracing method that shows the detailed morphology of neurons, their axons and terminals: immunohistochemical localization of an axonally transported plant lectin, Phaseolus vulgaris leucoagglutinin (PHA-L). Brain Res. 290 (2), 219–238. Gerfen, C.R., Sawchenko, P.E., 1985. A method for anterograde axonal tracing of chemically specified circuits in the central nervous system: combined Phaseolus vulgaris–leucoagglutinin (PHA-L) tract tracing and immunohistochemistry. Brain Res. 343 (1), 144–150. Glickstein, M., 2006. Golgi and Cajal: the neuron doctrine and the 100th anniversary of the 1906 Nobel Prize. Curr. Biol. 16 (5), R147–R151. Godschalk, M., Lemon, R.N., Kuypers, H.G., Ronday, H.K., 1984. Cortical afferents and efferents of monkey postarcuate area: an anatomical and electrophysiological study. Exp. Brain Res. 56 (3), 410–424. Gonatas, N.K., Harper, C., Mizutani, T., Gonatas, J.O., 1979. Superior sensitivity of conjugates of horseradish peroxidase with wheat germ agglutinin for studies of retrograde axonal transport. J. Histochem. Cytochem. 27 (3), 728–734. Graf, W., Gerrits, N., Yatim-Dhiba, N., Ugolini, G., 2002. Mapping the oculomotor system: the power of transneuronal labelling with rabies virus. Eur. J. Neurosci. 15 (9), 1557–1562. Grafstein, B., Forman, D.S., 1980. Intracellular transport in neurons. Physiol. Rev. 60 (4), 1167–1283. Grantyn, A., Brandi, A.M., Dubayle, D., Graf, W., Ugolini, G., Hadjidimitrakis, K., Moschovakis, A., 2002. Density gradients of trans-synaptically labeled collicular neurons after injections of rabies virus in the lateral rectus muscle of the rhesus monkey. J. Comp. Neurol. 451 (4), 346–361. Harrison, P.J., Hultborn, H., Jankowska, E., Katz, R., Storai, B., Zytnicki, D., 1984. Labelling of interneurones by retrograde transsynaptic transport of horseradish peroxidase from motoneurones in rats and cats. Neurosci. Lett. 45 (1), 15–19. He, S.Q., Dum, R.P., Strick, P.L., 1993. Topographic organization of corticospinal projections from the frontal lobe: motor areas on the lateral surface of the hemisphere. J. Neurosci. 13 (3), 952–980. Heimer, L., RoBards, M., 1981. Neuroanatomical Tract Tracing Methods. Plenum Press, New York. Holodny, A.I., Gor, D.M., Watts, R., Gutin, P.H., Ulug, A.M., 2005. Diffusion-tensor MR tractography of somatotopic organization of corticospinal tracts in the internal capsule: initial anatomic results in contradistinction to prior reports. Radiology 234, 649–653. Hoover, J.E., Strick, P.L., 1993. Multiple output channels in the basal ganglia. Science 259 (5096), 819–821. Horn, A.K., Buttner-Ennever, J.A., 1990. The time course of retrograde transsynaptic transport of tetanus toxin fragment C in the oculomotor system of the rabbit after injection into extraocular eye muscles. Exp. Brain Res. 81 (2), 353–362. Horn, A.K., Buttner-Ennever, J.A., Suzuki, Y., Henn, V., 1995. Histological identification of premotor neurons for horizontal saccades in monkey and man by parvalbumin immunostaining. J. Comp. Neurol. 359 (2), 350–363. Horton, J.C., Hocking, D.R., 1996. Anatomical demonstration of ocular dominance columns in striate cortex of the squirrel monkey. J. Neurosci. 16 (17), 5510–5522.


10017-JOHANSEN-BERG-9780123964601


396


Huisman, A.M., Kuypers, H.G., Verburgh, C.A., 1982. Differences in collateralization of the descending spinal pathways from red nucleus and other brain stem cell groups in cat and monkey. Prog. Brain Res. 57, 185–217. Innocenti, G.M., Clarke, S., Kraftsik, R., 1986. Interchange of callosal and association projections in the developing visual cortex. J. Neurosci. 6 (5), 1384–1409. Jacob, Y., Badrane, H., Ceccaldi, P.E., Tordo, N., 2000. Cytoplasmic dynein LC8 interacts with lyssavirus phosphoprotein. J. Virol. 74 (21), 10217–10222. Jeppesen, G.M., Hoerber, J.K., 2012. The mechanical properties of kinesin-1: a holistic approach. Biochem. Soc. Trans. 40 (2), 438–443. Johnson, T.N., Rosvold, H.E., Mishkin, M., 1968. Projections from behaviorally-defined sectors of the prefrontal cortex to the basal ganglia, septum, and diencephalon of the monkey. Exp. Neurol. 21 (1), 20–34. Jones, E.G., Powell, T.P., 1970. An anatomical study of converging sensory pathways within the cerebral cortex of the monkey. Brain 93 (4), 793–820. Jorritsma-Byham, B., Witter, M.P., Wouterlood, F.G., 1994. Combined anterograde tracing with biotinylated dextran-amine, retrograde tracing with fast blue and intracellular filling of neurons with lucifer yellow: an electron microscopic method. J. Neurosci. Methods 52 (2), 153–160. Kajiwara, R., Wouterlood, F.G., Sah, A., Boekel, A.J., Baks-te Bulte, L.T., Witter, M.P., 2008. Convergence of entorhinal and CA3 inputs onto pyramidal neurons and interneurons in hippocampal area CA1–an anatomical study in the rat. Hippocampus 18 (3), 266–280. Keizer, K., Kuypers, H.G., 1989. Distribution of corticospinal neurons with collaterals to the lower brain stem reticular formation in monkey (Macaca fascicularis). Exp. Brain Res. 74 (2), 311–318. Keizer, K., Kuypers, H.G., Huisman, A.M., Dann, O., 1983. Diamidino yellow dihydrochloride (DY:2HCl); a new fluorescent retrograde neuronal tracer, which migrates only very slowly out of the cell. Exp. Brain Res. 51 (2), 179–191. Kelly, R.M., Strick, P.L., 2000. Rabies as a transneuronal tracer of circuits in the central nervous system. J. Neurosci. Methods 103 (1), 63–71. Kelly, R.M., Strick, P.L., 2003. Cerebellar loops with motor cortex and prefrontal cortex of a nonhuman primate. J. Neurosci. 23 (23), 8432–8444. Kim, J.S., Kim, J., 2005. Pure midbrain infarction: clinical, radiologic, and pathophysiologic findings. Neurology 64 (7), 1227–1232. Kobbert, C., Apps, R., Bechmann, I., Lanciego, J.L., Mey, J., Thanos, S., 2000. Current concepts in neuroanatomical tracing. Prog. Neurobiol. 62 (4), 327–351. Kreig, W.J.S., 1954. Connections of the Cerebral Cortex. Thomas, Springfield, IL. Kristensson, K., Olsson, Y., 1971. Retrograde axonal transport of protein. Brain Res. 29 (2), 363–365. Kuypers, H.G.J.M., 1981. Anatomy of the descending pathways. In: Motor Control, pt, I., Brooks, V.B. (Eds.), Handbook of Physiology, Sec. I, The Nervous System, vol. II. American Physiological Society, Bethesda, MD, pp. 567–666. Kuypers, H.G.J.M., Huisman, A.M., 1984. Fluorescent neuronal tracers. In: Fedorff, S. (Ed.), Labeling Methods Applicable to the Study of Neuronal Pathways, Advances in Cellular Neurobiology, Vol. 5. Academic Press, New York, pp. 307–334. Kuypers, H.G., Ugolini, G., 1990. Viruses as transneuronal tracers. Trends. Neurosci. 13 (2), 71–75. Kuypers, H.G., Szwarcbart, M.K., Mishkin, M., Rosvold, H.E., 1965. Occipitotemporal corticocortical connections in the rhesus monkey. Exp. Neurol. 11, 245–262.

Kuypers, H.G.J.M., Catsman-Berrevoets, C.E., Padt, R.E., 1977. Retrograde axonal transport of fluorescent substances in the rat’s forebrain. Neurosci. Lett. 6, 127–135. Kuypers, H.G.J.M., Bentivoglio, M., Van der Kooy, D., CatsmanBerrevoets, C.E., 1979. Retrograde transport of bisbenzimide and propidium iodide through axons to their parent cell bodies. Neurosci. Lett. 12, 1–7. Kuypers, H.G., Bentivoglio, M., Catsman-Berrevoets, C.E., Bharos, A.T., 1980. Double retrograde neuronal labeling through divergent axon collaterals, using two fluorescent tracers with the same excitation wavelength which label different features of the cell. Exp. Brain Res. 40 (4), 383–392. Lacroix, S., Havton, L.A., McKay, H., Yang, H., Brant, A., Roberts, J., Tuszynski, M.H., 2004. Bilateral corticospinal projections arise from each motor cortex in macaque monkey: a quantitative study. J. Comp. Neurol. 473, 147–161. Lafon, M., 2005. Rabies virus receptors. J. Neurovirol. 11 (1), 82–87. Lanciego, J.L., Wouterlood, F.G., 1994. Dual anterograde axonal tracing with Phaseolus vulgaris leucoagglutinin (PHA-L) and biotinylated dextran amine (BDA). Neurosci. Prot. 94–050–06. Lanciego, J.L., Wouterlood, F.G., 2006. Multiple neuroanatomical tracttracing: approaches for multiple tract tracing. In: Zaborszky, L., Wouterlood, F.G., Lanciego, J.L. (Eds.), Neuroanatomical TractTracing 3: Molecules, Neurons, and Systems. Springer, New York, pp. 336–365. Lanciego, J.L., Wouterlood, F.G., 2011. A half century of experimental neuroanatomical tracing. J. Chem. Neuroanat. 42 (3), 157–183. Lanciego, J.L., Wouterlood, F.G., Erro, E., Gimenez-Amaya, J.M., 1998. Multiple axonal tracing: simultaneous detection of three tracers in the same section. Histochem. Cell. Biol. 110 (5), 509–515. Lang, C.E., Schieber, M.H., 2003. Differential impairment of individuated finger movements in humans after damage to the motor cortex or the corticospinal tract. J. Neurophysiol. 90 (2), 1160–1170. LaVail, J.H., LaVail, M.M., 1972. Retrograde axonal transport in the central nervous system. Science 176 (42), 1416–1417. Loewy, A.D., 1995. Pseudorabies virus: a transneuronal tracer for neuroanatomical studies. In: Keplitt, M., Lowey, A.D. (Eds.), Viral Vectors. Academic Press, New York, pp. 349–366. Luppi, P.H., Fort, P., Jouvet, M., 1990. Iontophoretic application of unconjugated cholera toxin B subunit (CTb) combined with immunohistochemistry of neurochemical substances: a method for transmitter identification of retrogradely labeled neurons. Brain Res. 534 (1–2), 209–224. Lyckman, A.W., Fan, G., Rios, M., Jaenisch, R., Sur, M., 2005. Normal eye-specific patterning of retinal inputs to murine subcortical visual nuclei in the absence of brain-derived neurotrophic factor. Vis. Neurosci. 22 (1), 27–36. Manni, E., Troiani, D., 2005. Vittorio Marchi’s method of staining degenerating nervous fibers and the tragedy of his life. Arch. Neurol. 62 (2), 321–325. Marchi, V., Algeri, G., 1885. Sulle degenerazioni discendenti consecutive a lesioni sperimentali in diverse zone della corteccia cerebrale. Riv. Sper. Freniatria. Med. Legal. 11, 492–494. Mauro, A., Germano, I., Giaccone, G., Giordana, M.T., Schiffer, D., 1985. 1-Naphthol basic dye (1-NBD). An alternative to diaminobenzidine (DAB) in immunoperoxidase techniques. Histochemistry 83 (2), 97–102. McNeal, D.W., Ge, J., Herrick, J.L., Stilwell-Morecraft, K.S., Morecraft, R.J., 2008. Characterization of some morphological parameters of orbicularis oculi motor neurons in the monkey. Neuroscience 151 (1), 12–27. McNeal, D.W., Darling, W.G., Ge, J., Stilwell-Morecraft, K.S., Solon, K.M., Hynes, S.M., Pizzimenti, M.A., Rotellla, D.L., Vanadurongvan, T., Morecraft, R.J., 2010. Selective long-term reorganization of the corticospinal projection from the


10017-JOHANSEN-BERG-9780123964601


REFERENCES

supplementary motor cortex following recovery from lateral motor cortex injury. J. Comp. Neurol. 518, 586–621. Meissirel, C., Dehay, C., Berland, M., Kennedy, H., 1991. Segregation of callosal and association pathways during development in the visual cortex of the primate. J. Neurosci. 11 (11), 3297–3316. Mesulam, M.M., 1978. Tetramethyl benzidine for horseradish peroxidase neurohistochemistry: a non-carcinogenic blue reaction product with superior sensitivity for visualizing neural afferents and efferents. J. Histochem. Cytochem. 26 (2), 106–117. Mesulam, M.M., 1982. Tracing Neural Connections with Horseradish Peroxidase. Wiley, New York. Mettler, F.A., 1935a. Corticofugal fiber connections of the cerebral cortex of the Macaca mulatta. The occipital region. J. Comp. Neurol. 61, 221–256. Mettler, F.A., 1935b. Corticofugal fiber connections of the cerebral cortex of the Macaca mulatta. The frontal region. J. Comp. Neurol. 61, 509–542. Mettler, F.A., 1935c. Corticofugal fiber connections of the cerebral cortex of the Macaca mulatta. The parietal region. J. Comp. Neurol. 62, 263–291. Mettler, F.A., 1935d. Corticofugal fiber connections of the cerebral cortex of the Macaca mulatta. The temporal region. J. Comp. Neurol. 63, 25–47. Meyers, R., 1951. The human frontocorticopontine tract: functional inconsequence of its surgical interruption. Neurology 1 (5), 341–356. Middleton, F.A., Strick, P.L., 1994. Anatomical evidence for cerebellar and basal ganglia involvement in higher cognitive function. Science 266 (5184), 458–461. Miyachi, S., Lu, X., Imanishi, M., Sawada, K., Nambu, A., Takada, M., 2006. Somatotopically arranged inputs from putamen and subthalamic nucleus to primary motor cortex. Neurosci. Res. 56 (3), 300–308. Morcuende, S., Delgado-Garcia, J.M., Ugolini, G., 2002. Neuronal premotor networks involved in eyelid responses: retrograde transneuronal tracing with rabies virus from the orbicularis oculi muscle in the rat. J. Neurosci. 22 (20), 8808–8818. Morecraft, R.J., Guela, C., Mesulam, M.M., 1992. Cytoarchitecture and neural afferents of orbitofrontal cortex in brain of the monkey. J. Comp. Neurol. 323, 341–358. Morecraft, R.J., Louie, J.L., Herrick, J.L., Stilwell-Morecraft, K.S., 2001. Cortical innervation of the facial nucleus in the non-human primate: a new interpretation of the effects of stroke and related subtotal brain trauma on the muscles of facial expression. Brain 124 (Pt 1), 176–208. Morecraft, R.J., Herrick, J.L., Stilwell-Morecraft, K.S., Louie, J.L., Schroeder, C.M., Ottenbacher, J.G., Schoolfield, M.W., 2002. Localization of arm representation in the corona radiata and internal capsule in the non-human primate. Brain 125 (Pt 1), 176–198. Morecraft, R.J., Cipolloni, P.B., Stilwell-Morecraft, K.S., Gedney, M.T., Pandya, D.N., 2004. Cytoarchitecture and cortical connections of the posterior cingulate and adjacent somatosensory fields in the rhesus monkey. J. Comp. Neurol. 469, 37–69. Morecraft, R.J., McNeal, D.W., Stilwell-Morecraft, K.S., Dvanajscak, Z., Ge, J., Schneider, P., 2007a. Localization of arm representation in the cerebral peduncle of the non-human primate. J. Comp. Neurol. 504 (2), 149–167. Morecraft, R.J., McNeal, D.W., Stilwell-Morecraft, K.S., Gedney, M., Ge, J., Schroeder, C.M., Van Hoesen, G.W., 2007b. Amygdala interconnections with the cingulate motor cortex in the rhesus monkey. J. Comp. Neurol. 500 (1), 134–165. Morecraft, Rj, Stilwell-Morecraft, K.S., Cipolloni, P.B., Ge, J., McNeal, D.W., Pandya, D.N., 2012. Cytoarchitecture and cortical connections of the anterior cingulate and adjacent somatomotor fields in the rhesus monkey. Brain. Res. Bull. 87, 457–497.

397

Moschovakis, A.K., Gregoriou, G.G., Ugolini, G., Doldan, M., Graf, W., Guldin, W., Hadjidimitrakis, K., Savaki, H.E., 2004. Oculomotor areas of the primate frontal lobes: a transneuronal transfer of rabies virus and [14C]-2-deoxyglucose functional imaging study. J. Neurosci. 24 (25), 5726–5740. Myers, R.E., 1962. Commissural connections between occipital lobes of the monkey. J. Comp. Neurol. 118, 1–16. Nance, D.M., Burns, J., 1990. Fluorescent dextrans as sensitive anterograde neuroanatomical tracers: applications and pitfalls. Brain. Res. Bull. 25, 139–145. Nauta, W.J., 1964. Some efferent connections of the prefrontal cortex in the monkey. In: Waren, J.M., Akert, K. (Eds.), The Frontal Granular Cortex and Behavior. McGraw-Hill, New York, pp. 397–409. Nauta, W.J., 1972. Neural associations of the frontal cortex. Acta. Neurobiol. Exp. (Wars) 32 (2), 125–140. Nauta, W.J., 1993. Some early travails of tracing axonal pathways in the brain. J. Neurosci. 13 (4), 1337–1345. Nauta, W.J., Gygax, P.A., 1951. Silver impregnation of degenerating axon terminals in the central nervous system: (1) Technic. (2) Chemical notes. Stain. Technol. 26 (1), 5–11. Nauta, W.J., Gygax, P.A., 1954. Silver impregnation of degenerating axons in the central nervous system: a modified technic. Stain. Technol. 29 (2), 91–93. Newton, J.M., Ward, N.S., Parker, G.J., Deichmann, R., Alexander, D.C., Friston, K.J., Frackowiak, R.S., 2006. Non-invasive mapping of corticofugal fibers from multiple motor areas– relevance to stroke recovery. Brain 129, 1844–1858. Ochs, S., 1979. The early history of material transport in nerve. Physiologist 22 (5), 16–19. Ochs, S., 2004. A History of Nerve Functions: From Animal Spirits to Molecular Mechanisms. Cambridge University Press, Cambridge. Orioli, P.J., Strick, P.L., 1989. Cerebellar connections with the motor cortex and the arcuate premotor area: an analysis employing retrograde transneuronal transport of WGA-HRP. J. Comp. Neurol. 288 (4), 612–626. Oztas, E., 2003. Neuronal tracing. Neuroanatomy 2, 2–5. Pandya, D.N., Kuypers, H.G., 1969. Cortico-cortical connections in the rhesus monkey. Brain Res. 13 (1), 13–36. Pandya, D.N., Karol, E.A., Heilbronn, D., 1971. The topographical distribution of interhemispheric projections in the corpus callosum of the rhesus monkey. Brain Res. 32, 31–43. Perreault, M.C., Bernier, A.P., Renaud, J.S., Roux, S., Glover, J.C., 2006. C fragment of tetanus toxin hybrid proteins evaluated for musclespecific transsynaptic mapping of spinal motor circuitry in the newborn mouse. Neuroscience 141 (2), 803–816. Pinault, D., 1996. A novel single-cell staining procedure performed in vivo under electrophysiological control: morpho-functional features of juxtacellularly labeled thalamic cells and other central neurons with biocytin or Neurobiotin. J. Neurosci. Methods 65 (2), 113–136. Porter, J.D., Guthrie, B.L., Sparks, D.L., 1985. Selective retrograde transneuronal transport of wheat germ agglutinin-conjugated horseradish peroxidase in the oculomotor system. Exp. Brain Res. 57 (2), 411–416. Prevosto, V., Graf, W., Ugolini, G., 2009. Posterior parietal cortex areas MIP and LIPv receive eye position and velocity inputs via ascending preposito-thalamo-cortical pathways. Eur. J. Neurosci. 30 (6), 1151–1161. Prevosto, V., Graf, W., Ugolini, G., 2010. Cerebellar inputs to intraparietal cortex areas LIP and MIP: functional frameworks for adaptive control of eye movements, reaching, and arm/eye/head movement coordination. Cereb. Cortex. 20 (1), 214–228. Prevosto, V., Graf, W., Ugolini, G., 2011. Proprioceptive pathways to posterior parietal areas MIP and LIPv from the dorsal column nuclei and the postcentral somatosensory cortex. Eur. J. Neurosci. 33 (3), 444–460.


10017-JOHANSEN-BERG-9780123964601


398 ½AU1


Raghavan, P., Krakauer, J.W., Gordon, A.M., 2006. Impaired anticipatory control of fingertip forces in patients with a pure motor or sensorimotor lucanar syndrome. Brain 129, 1415–1425. Rathelot, J.A., Strick, P.L., 2006. Muscle representation in the macaque motor cortex: an anatomical perspective. Proc. Natl. Acad. Sci. U. S. A. 103 (21), 8257–8262. Reiner, A., Anderson, K.D., 1993. Co-occurrence of gamma-aminobutyric acid, parvalbumin and the neurotensin-related neuropeptide LANT6 in pallidal, nigral and striatal neurons in pigeons and monkeys. Brain Res. 624 (1–2), 317–325. Reiner, A., Veenman, C.L., Medina, L., Jiao, Y., Del Mar, N., Honig, M.G., 2000. Pathway tracing using biotinylated dextran amines. J. Neurosci. Methods 103 (1), 23–37. Robertson, B., Arvidsson, J., 1985. Transganglionic transport of wheat germ agglutinin-HRP and choleragenoid-HRP in rat trigeminal primary sensory neurons. Brain Res. 348 (1), 44–51. Robertson, B., Grant, G., 1985. A comparison between wheat germ agglutinin and choleragenoid–horseradish peroxidase as anterogradely transported markers in central branches of primary sensory neurones in the rat with some observations in the cat. Neuroscience 14 (3), 895–905. Rosene, D.L., Roy, N.J., Davis, B.J., 1986. A cryoprotection method that facilitates cutting frozen sections of whole monkey brains for histological and histochemical processing without freezing artifact. J. Histochem. Cytochem. 34 (10), 1301–1315. Rosenzweig, E.S., Courtine, G., Jindrich, D.L., Brock, J.H., Ferguson, A.R., Strand, S.C., Nout, Y.S., Roy, R.R., Miller, D.M., Beattie, M.S., Havton, L.A., Bresnahan, J.C., Edgerton, V.R., Tuszynski, M.H., 2010. Extensive spontaneous plasticity of corticospinal projections after primate spinal cord injury. Nature. Neurosci. 13 (12), 1505–1510. Rosina, A., 1982. Rapid anterograde movement of the fluorescent tracer Fast Blue: a new method for tracing central connections. Neurosci. Lett. 33 (3), 217–221. Rosoklija, G., Mancevski, B., Ilievski, B., Perera, T., Lisanby, S.H., Coplan, J.D., Duma, A., Serafimova, T., Dwork, A.J., 2003. Optimization of Golgi methods for impregnation of brain tissue from humans and monkeys. J. Neurosci. Methods 131 (1–2), 1–7. Rotto-Percelay, D.M., Wheeler, J.G., Osorio, F.A., Platt, K.B., Loewy, A.D., 1992. Transneuronal labeling of spinal interneurons and sympathetic preganglionic neurons after pseudorabies virus injections in the rat medial gastrocnemius muscle. Brain Res. 574 (1–2), 291–306. Saleem, K.S., Pauls, J.M., Augath, M., Trinath, T., Prause, B.A., Hashikawa, T., Logothetis, N.K., 2002. Magnetic resonance imaging of neuronal connections in the macaque monkey. Neuron 34 (5), 685–700. Salin, P., Lo´pez, I.P., Kachidian, P., Barroso-Chinea, P., Rico, A.J., Go´mez-Bautista, V., Coulon, P., Kerkerian-Le Goff, L., Lanciego, J.L., 2009. Changes to interneuron-driven striatal microcircuits in a rat model of Parkinson’s disease. Neurobiol. Dis. 34 (3), 545–552. Schaechter, J.D., Perdue, K.L., Wang, R., 2008. Structural damage to the corticospinal tract correlates with bilateral sensorimotor cortex reorganization in stroke patients. NeuroImage 39 (3), 1370–1382. Schiemanck, S.K., Kwakkel, G., Post, M.W.M., Kappelle, L.J., Prevo, A.J.H., 2008. Impact of internal capsule lesions on outcome of motor hand function at one year post-stroke. J. Rehabil. Med. 40, 96–101. Schmahmann, J.D., Pandya, D.N., 2006. Fiber Pathways of the Brain. Oxford University Press, Oxford. Schmued, L.C., Fallon, J.H., 1986. Fluoro-Gold: a new fluorescent retrograde axonal tracer with numerous unique properties. Brain Res. 377 (1), 147–154.

Schmued, L., Kyriakidis, K., Heimer, L., 1990. In vivo anterograde and retrograde axonal transport of the fluorescent rhodamine-dextranamine, Fluoro-Ruby, within the CNS. Brain Res. 526 (1), 127–134. Schnell, M.J., McGettigan, J.P., Wirblich, C., Papaneri, A., 2010. The cell biology of rabies virus: using stealth to reach the brain. Nat. Rev. Microbiol. 8 (1), 51–61. Schwab, M.E., Javoy-Agid, F., Agid, Y., 1978. Labeled wheat germ agglutinin (WGA) as a new, highly sensitive retrograde tracer in the rat brain hippocampal system. Brain Res. 152 (1), 145–150. Shelton, F.N., Reding, M.J., 2001. Effect of lesion location on upper limb motor recovery after stroke. Stroke 32 (1), 107–112. Singer, C., 1952. Vesalius on the Human Brain. Oxford University Press, London. Singer, W., Hollander, H., Vanegas, H., 1977. Decreased peroxidase labeling of lateral geniculate neurons following deafferentation. Brain Res. 120 (1), 133–137. Spiga, S., Acquas, E., Puddu, M.C., Mulas, G., Lintas, A., Diana, M., 2011. Simultaneous Golgi-Cox and immunofluorescence using confocal microscopy. Brain. Struct. Funct. 216 (3), 171–182. Sullivan, E.V., Zahr, N.M., Rohlfing, T., Pfefferbaum, A., 2010. Fiber tracking functionally distinct components of the internal capsule. Neuropsychologia 48 (14), 4155–4163. Swanson, L.W., 2000. A history of neuroanatomical mapping. In: Toga, A.W., Mazziotta, J.C. (Eds.), Brain Mapping: The Applications. Academic Press, San Diego, pp. 77–109. Tang, Y., Rampin, O., Giuliano, F., Ugolini, G., 1999. Spinal and brain circuits to motoneurons of the bulbospongiosus muscle: retrograde transneuronal tracing with rabies virus. J. Comp. Neurol. 414 (2), 167–192. Thomas, M.A., Lemmer, B., 2005. HistoGreen: a new alternative to 3,30 diaminobenzidine-tetrahydrochloride-dihydrate (DAB) as a peroxidase substrate in immunohistochemistry? Brain Res. Brain Res. Protoc. 14 (2), 107–118. Trojanowski, J.Q., 1983. Native and derivatized lectins for in vivo studies of neuronal connectivity and neuronal cell biology. J. Neurosci. Methods 9 (3), 185–204. Trojanowski, J.Q., Gonatas, J.O., Gonatas, N.K., 1981. Conjugates of horseradish peroxidase (HRP) with cholera toxin and wheat germ agglutinin are superior to free HRP as orthogradely transported markers. Brain Res. 223 (2), 381–385. Trojanowski, J.Q., Gonatas, J.O., Gonatas, N.K., 1982. Horseradish peroxidase (HRP) conjugates of cholera toxin and lectins are more sensitive retrogradely transported markers than free HRP. Brain Res. 231 (1), 33–50. Turner, P.T., 1977. Effect of pentobarbital on uptake of horseradish peroxidase by rabbit cortical synapses. Exp. Neurol. 54 (1), 24–32. Ugolini, G., 1992. Transneuronal transfer of herpes simplex virus type 1 (HSV 1) from mixed limb nerves to the CNS. I. Sequence of transfer from sensory, motor, and sympathetic nerve fibres to the spinal cord. J. Comp. Neurol. 326 (4), 527–548. Ugolini, G., 1995a. Specificity of rabies virus as a transneuronal tracer of motor networks: transfer from hypoglossal motoneurons to connected second-order and higher order central nervous system cell groups. J. Comp. Neurol. 356 (3), 457–480. Ugolini, G., 1995b. Transneuronal tracing with alpha-herpesviruses: a review of the methodology. In: Keplitt, M., Loewy, A.D. (Eds.), Viral Vectors: Gene Therapy and Neuroscience Applications. Academic Press, New York, pp. 293–317. Ugolini, G., 1996. Tracing connected neurons with Herpes Simplex Virus type 1. In: Lowenstein, P.R., Enquist, L.W. (Eds.), Protocols for Gene Transfer in Neuroscience: Towards Gene Therapy of Neurological Disorders. John Wiley & Sons, Chichester, pp. 349–364. Ugolini, G., 2008. Use of rabies virus as a transneuronal tracer of neuronal connections: implications for the understanding of rabies pathogenesis. Dev. Biol. (Basel) 131, 493–506.


10017-JOHANSEN-BERG-9780123964601


FURTHER READING

Ugolini, G., 2010. Advances in viral transneuronal tracing. J. Neurosci. Methods 194 (1), 2–20. Ugolini, G., 2011. Rabies virus as a transneuronal tracer of neuronal connections. Adv. Virus. Res. 79, 165–202. Ugolini, G., Kuypers, H.G., 1986. Collaterals of corticospinal and pyramidal fibres to the pontine grey demonstrated by a new application of the fluorescent fibre labelling technique. Brain Res. 365 (2), 211–227. Ugolini, G., Kuypers, H.G., Simmons, A., 1987. Retrograde transneuronal transfer of herpes simplex virus type 1 (HSV 1) from motoneurones. Brain Res. 422 (2), 242–256. Ugolini, G., Kuypers, H.G., Strick, P.L., 1989. Transneuronal transfer of herpes virus from peripheral nerves to cortex and brainstem. Science 243 (4887), 89–91. Ugolini, G., Klam, F., Doldan Dans, M., Dubayle, D., Brandi, A.M., Buttner-Ennever, J., Graf, W., 2006. Horizontal eye movement networks in primates as revealed by retrograde transneuronal transfer of rabies virus: differences in monosynaptic input to “slow” and “fast” abducens motoneurons. J. Comp. Neurol. 498 (6), 762–785. Van Hoesen, G.W., 1993. The modern concept of association cortex. Curr. Opin. Neurobiol. 3 (2), 150–154. Van Hoesen, G.W., Pandya, D.N., Butters, N., 1972. Cortical afferents to the entorhinal cortex of the rhesus monkey. Science 175 (4029), 1471–1473. Veenman, C.L., Reiner, A., Honig, M.G., 1992. Biotinylated dextran amine as an anterograde tracer for single-and double-labeling studies. J. Neurosci. Methods 41 (3), 239–254. Walker, A.E., 1952. Cerebral pedunculotomy for the relief of involuntary movements. II. Parkinsonian tremor. J. Nerv. Ment. Dis. 116, 766–775. Waller, A., 1850. Experiments on the section of the glossopharyngeal and hypoglossal nerves of the frog and observations of the alterations produced thereby in the structures of their primitive fibers. Phil. Trans. R. Soc. 140, 423–429. Wan, X.C., Trojanowski, J.Q., Gonatas, J.O., 1982. Cholera toxin and wheat germ agglutinin conjugates as neuroanatomical probes: their uptake and clearance, transganglionic and retrograde transport and sensitivity. Brain Res. 243 (2), 215–224. Warr, W.B., de Olmos, J.S., Heimer, L., 1981. Horseradish peroxidase: the basic procedure. In: Heimer, L., RoBards, M. (Eds.), Neuroanatomical Tract-Tracing Methods. Plenum, New York, pp. 207–262. Wenzelburger, R., Kopper, F., Frenzel, A., Stolze, H., Klebe, S., Brossmann, A., Kuhtz-Buschbeck, J., Golge, M., Illert, M., Deuschl, G., 2005. Hand coordination following capsular stroke. Brain 128 (Pt 1), 64–74. Wiesel, T.N., Hubel, D.H., Lam, D.M., 1974. Autoradiographic demonstration of ocular-dominance columns in the monkey striate cortex by means of transneuronal transport. Brain Res. 79 (2), 273–279. Wouterlood, F.G., 1991. Innervation of entorhinal principal cells by neurons of the nucleus reuniens thalami. Anterograde PHA-L tracing combined with retrograde fluorescent tracing and intracellular injection with lucifer yellow in the rat. Eur. J. Neurosci. 3 (7), 641–647. Wouterlood, F.G., Jorritsma-Byham, B., 1993. The anterograde neuroanatomical tracer biotinylated dextran-amine: comparison with the tracer Phaseolus vulgaris-leucoagglutinin in preparations for electron microscopy. J. Neurosci. Methods 48 (1–2), 75–87.

399

Wouterlood, F.G., Bol, J.G., Steinbusch, H.W., 1987. Double-label immunocytochemistry: combination of anterograde neuroanatomical tracing with Phaseolus vulgaris leucoagglutinin and enzyme immunocytochemistry of target neurons. J. Histochem. Cytochem. 35 (8), 817–823. Wouterlood, F.G., Goede, P.H., Arts, M.P., Groenewegen, H.J., 1992. Simultaneous characterization of efferent and afferent connectivity, neuroactive substances, and morphology of neurons. J. Histochem. Cytochem. 40 (4), 457–465. Wouterlood, F.G., Goede, P.H., Jorritsma-Byham, B., Witter, M.P., 1997. Neuroanatomical tracing based on cellular transport: horseradish peroxidase, Phaseolus vulgaris-leucoagglutinin, and biotinylated dextran amine. In: O’Mara, S. (Ed.), Neurobiology Labfax. Academic Press, London, pp. 33–46. Wouterlood, F.G., Bockers, T., Witter, M.P., 2003. Synaptic contacts between identified neurons visualized in the confocal laser scanning microscope. Neuroanatomical tracing combined with immunofluorescence detection of post-synaptic density proteins and target neuron-markers. J. Neurosci. Methods 128 (1–2), 129–142. Wouterlood, F.G., Boekel, A.J., Meijer, G.A., Belien, J.A., 2007. Computer-assisted estimation in the CNS of 3D multimarker “overlap” or “touch” at the level of individual nerve endings: a confocal laser scanning microscope application. J. Neurosci. Res. 85 (6), 1215–1228. Wouterlood, F.G., Aliane, V., Boekel, A.J., Hur, E.E., Zaborszky, L., Barroso-Chinea, P., Hartig, W., Lanciego, J.L., Witter, M.P., 2008a. Origin of calretinin-containing, vesicular glutamate transporter 2coexpressing fiber terminals in the entorhinal cortex of the rat. J. Comp. Neurol. 506 (2), 359–370. Wouterlood, F.G., Boekel, A.J., Kajiwara, R., Belien, J.A., 2008b. Counting contacts between neurons in 3D in confocal laser scanning images. J. Neurosci. Methods 171 (2), 296–308. Yoshino-Saito, K., Nishimura, Y., Oishi, T., Isa, T., 2010. Quantitative inter-segmental and inter-laminar comparison of corticospinal projections from the forelimb area of the primary motor cortex of macaque monkeys. Neuroscience 171, 1164–1179. Zemanick, M.C., Strick, P.L., Dix, R.D., 1991. Direction of transneuronal transport of herpes simplex virus 1 in the primate motor system is strain-dependent. Proc. Natl. Acad. Sci. U. S. A. 88 (18), 8048–8051. Zhou, M., Grofova, I., 1995. The use of peroxidase substrate Vector VIP in electron microscopic single and double antigen localization. J. Neurosci. Methods 62 (1–2), 149–158. Zimber, M.S., 2001. Vesalius’ Fabrica: the marriage of art and anatomy. Arch. Facial. Plast. Surg. 3, 220–221.

Further Reading Bolam, J.P., 1992. Experimental Neuroanatomy. A Practical Approach. Oxford University Press, Oxford. Heimer, L., RoBards, M., 1981. Neuroanatomical Tract Tracing Methods. Plenum Press, New York. Heimer, L., Zaborszky, L., 1989. Neuroanatomical Tract Tracing Methods 2: Recent Progress. Plenum Press, New York. Zaborszky, L., Wouterlood, F.G., Lanciego, J.L., 2006. Neuroanatomical Tract-Tracing 3: Molecules, Neurons, and Systems. Springer, New York.


10017-JOHANSEN-BERG-9780123964601

JOHANSEN-BERG: 17 Non-Print Items

Abstract: Investigating the organization of nerve pathways is of fundamental interest in neurology and the neurosciences. In this chapter we draw attention to landmark events that have contributed to the scientific evolution of neuron tract tracing and cover in detail contemporary methods employed in experimental animals to trace nerve pathways. The basic anatomical tract-tracing methods, including the cellular mechanisms underlying tract tracer uptake and axonal transport, are described. Specific techniques and experimental applications involving tracers transported retrogradely, anterogradely, and transneuronally are reviewed. Finally, examples of combined neuroanatomical tract-tracing applications are discussed and illustrated. Keywords: Afferent pathway, anterograde tracer, co-localization, efferent pathway, history of tract tracing, immunohistochemistry, neuroanatomy, neuronal tract tracer, retrograde tracer, transneuronal tract tracer, viral tract tracer.