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PHYSIOLOGIC RESPONSES TO INFLAMMATION IN ISOLATED EQUINE JOINTS

DISSERTATION

Presented in Partial Fulfilment of the Requirements for

the Degree of Doctor of Philosophy in the Graduate

School of The Ohio State University

B y

Joanne Hardy, D.V.M., M.S.

ÿ >{c ÿ ifc 3|c

The Ohio State University 1996

Dissertation Committee:

A.L. Bertone, Adviser

W.W. Muir S.E. Weisbrode A dviser T.M. O’Dorisio Department of Veterinary Clinical Sciences

D.N. Granger UMI Number: 9710576

This microform edition is protected against unauthorized copying under Title 17, United States Code.

UMI 300 North Zeeb Road Ann Arbor, MI 48103 ABSTRACT

Using this model, the local Starling’s forces in the isolated joint at isogravimetric state are characterized and alterations in Starling’s forces in response to hemodynamic manipulations are described. In a separate series of experiments, articular inflammation was induced in the isolated innervated or denervated joint organ and local hemodynamic, fluid exchanges, metabolic responses, alterations in permeability to small and large molecules, inflammatory mediator release, cellular events and articular cartilage metabolic alterations are described.

Two preliminary studies were performed in the development of the isolated joint model. The first studies was essential to gain a better understanding of the local biomechanical behavior of the fluid filled closed joint cavity. In this study, the investigation of pressure volume curves in the joint revealed a sigmoid relationship, with low compliance at normal subatmospheric pressures, and gradually increasing compliance at supraatmospheric pressures of up to 30 mmHg. Thereafter, compliance of the joint again decreased exponentially. This study also revealed an important function of synovial fluid to increase joint compliance and hysteresis therefore preventing collapse of the joint cavity and allow fluid exchanges at subatmospheric pressures. Furthermore this study emphasized the importance of joint angle on pressure-volume relationships. In the second preliminary study, a colored microsphere technique was validated for determination of blood flow to the synovial membrane and joint capsule. Furthermore, that study documented a significant decrease in synovial membrane blood flow down to 16% of baseline at intraarticular pressures of 30 mmHg or greater.

The development and physiologic responses of the isolated model were subsequently described. The extracorporeal pump perfused isolated joint unit developed in this model was successfully be maintained for 6 hours. Using this system, isogravimetric state was defined at a circuit arterial pressure of 133 mmHg and venous pressure of 10.5 mm Hg. Starling’s forces were defined for this system. Arterial and venous pressure manipulations revealed that increases in arterial pressure significantly raised transsynovial flow and synovial fluid production, when lAP was maintained at atmospheric pressure. A similar but much less important response was observed with increased venous pressure.

Acute inflammation was successfully induced in the isolated innervated or denervated joint preparation, using interleukin-IB. Acute inflammation induced a significant increase in oxygen consumption and extraction ratio in response to increased metabolic demand. Acute inflammation increased synovial membrane permeability to albumin, but denervation lowered permeability to large molecules (MW 144,(XX)). Inflammation significantly increased synovial fluid production and the permeability surface area product of the synovial membrane

Acute inflammation was associated with a significant cellular response characterized by neutrophilic leukocytosis in synovial fluid and a neutrophilic vasculitis in the synovial membrane. A significant increase in interleukin-6, prostaglandin 5% and substance P was

ui detected in synovial fluid collected from inflamed joints. Inflammation induced a significant degradative response in articular cartilage, even after this short term exposure to an inflammatory mediator.

The isolated joint model described in these studies provided new insights in our understanding of articular physiology, an allowed to study of all pathophysiologic processes involved in articular inflammation. This model should be useful to further our knowledge of the response of joint to various disease processes, and should be very suitable to study the pharmacokinetics and pharmacodynamics of systemic or intraarticular therapies.

IV ACKNOWLEDGMENTS

I express sincere appreciation to Dr Alicia L Bertone for her guidance and enthousiasm throughout this research, and Dr William W Muir for his great insight, ideas and support Thanks go to Dr Steve Weisbrode, for his enthousiasm and for teaching me to understand histopathology. Thanks go to the other members of my advisory committee.

Dr Tom O’Dorisio and Dr Neil Granger, for their time, suggestions and comments. The technical assistance of Mrs Kena Chapman, Mrs Robin Nakkula, and Ms Heather

Burkhardt is gratefully acknowledged. VITA

September 17th, 1957 ...... Bom- Montreal, Quebec, Canada

1982 ...... D.V.M., University of Montreal, Montreal, Quebec, Canada

1983 ...... 1.P.S.A.V., Internat de Perfectionnement en Sciences Appliquées Vétérinaires, University of Montreal, Montreal, Quebec, Canada

1989 ...... Residency, Equine Surgery, The Ohio State University, Columbus, Ohio

1991 Diplomate, American College of Veterinary Surgeons

PUBLICATIONS

Moore RM, Hance S, Hardy J. Colonic luminal pressure in horses with strangulating and nonstrangulating obstruction of the large colon. Vet Surg 1996; 25:134-141.

Hardv J. Bertone AL, Muir WW. Joint pressure influences synovial tissue blood flow as determined by colored microspheres. J Appl Physiol 1996; 80:1225-1232.

Beard WL, Hardy J. Techniques de fixation de fractures des incisives par cerclage interdentaire chez le cheval. Pratique Vétérinaire Equine 1995; 27:159-166.

Moore RM, Hardy J. Muir WW. Mural blood flow distribution in the large colon of horses during low-flow ischemia and reperfusion. Am J Vet Res 1995,56: 812-818.

Hardy J. Bertone AL, Muir WW. Pressure-volume relationships in the equine carpus. J Appl Physiol 1995, 78:1977-1984.

Hardy J. Bednarski RM, Biller DS. Effect of phenylephrine on hemodynamics and splenic dimensions in the horse. Am J Vet Res 1994,55: 1570-1578.

VI Carter BG, Schneider RK, Hardy J. Bramlage LR and Bertone AL. Assessment and treatment of equine humerai fractures: retrospective study of 54 cases (1972-1990). Equine VetJ 1993; 25; 3:203-207

Hardy J. Bertone AL. Surgery of the equine large and small colon. Compend Cont Educ Pract Vet 1992; 14(11): 1501-1506.

Hardy J. Stewart RH, Beard WL, Yvorchuk K. Complications of nasogastric intubation in the horse: 9 cases (1987-1989). J Am Vet Med Assoc 1992; 201 (3); 483-486.

Hardy J. Robertson JT, Reed SM. Pericardiectomy for treatment of constrictive pericarditis in a mare. Equine Vet J 1992; 24.

Giraudet A, Hardy J. Reparation chirurgicale des déchirures du col utérin chez la jument. Technique et nouvelle approche d'anesthesie locale. Pratique Vétérinaire Equine 1991; 23(l):65-70.

Hardy J. Marcoux M, Breton L. Clinical relevance of radiographie findings in the proximal sesamoid bones of two-year-old Standardbreds in their first year in training. J Am Vet Med Assoc 1991; 198(12):2089-2094.

Hardy J. Shiroma J. Temporomandibular luxation in a horse. J Am Vet Med Assoc 1991 ; 198(9): 1663-1664.

Hardy J. Upper airway obstmction in foals, weanlings and yearlings. Veterinary Clinics of North America:Equine Practice 1991;7(1):105-122.

Hardy J. Giraudet A. Revue clinique: Les affections des poches gutturales Clinical review: Diseases of the guttural pouches. Pratique Vétérinaire Equine 1990; 42(4):42-50.

Hardy J. Robertson JT, Wilkie DA. Ischemic optic neuropathy and blindness following arterial occlusion for treatment of guttural pouch mycosis in two horses. J Am Vet Med Assoc 1990(10):1631-1634.

Hardy J. Marone M. Luxation of the scapulohumeral joint in a foal. J Am Vet Med Assoc 1989; 195(12): 1773-1774.

Hardy J, Muir WW. Ventricular tachycardia associated with hyperkalemia in a mare, in ECG of the month. J Am Vet Med Assoc 1989; 194(3):356-357.

Hardv J. Marcoux M, Breton L. Prevalence and description of articular fragments of the fetlock joint in the Standardbred horse. Pratique Veterinaire Equine 1987; 19(4):27-32.

Hardv J. Marcoux M, Breton L. Prevalence and description of articular fragments of the fetlock joint in the Standardbred horse. Med Vet Quebec 1987;17(2):57-61.

Hardv J. Marcoux M, Breton L. Osteochondrosis-like lesion of the anconeal process in two horses. J Am Vet Med Assoc 1986; 189(7):802-803.

vu Hardy J. Marcoux M. Sacro-iliac arthrosis in the Standardbred Horse. Med Vet Quebec 1985; 15(4): 185-189.

Book chapters

Hardy J. Disorders of the proximal sesamoid bones. In Current Therapy in Equine Medicine, 4th ed. NE Robinson, editor. WB Saunders Co, Philadelphia. In press.

Hardv J. Chapter 38. Diseases of soft tissues, in The Horse: Diseases and clinical management. Kobluk CN, Ames TR, Geor RJ, editors. WB Saunders Co, Philadelphia, 1995, pp 791-814.

Hardy J. Sesamoiditis and suspensory desmitis. in Current Therapy in Equine Medicine, 3rd ed. NE Robinson, editor. WB Saunders Co, Philadelphia, 1992, pp 140-143.

Hardy J. Chapter 56. Surgical procedures involving the peripheral nerves, in Textbook of Equine Surgery. JA Auer editor. WB Saunders Co, Philadelphia, 1992, pp 580-585.

Proceedings and abstracts

-Repair of incisor fractures in the horse by interdental wiring. Ohio Veterinary Medical Association Annual Convention, Columbus, OH 1996.

- Blood flow, permeability and 02 metabolism of innervated or denervated isolated joints in an D-l model. Advances in Veterinary Research Day, The Ohio State University, Feb 1996.

- Blood flow, permeability and 02 metabolism of innervated or denervated isolated joints in an D-l model. Trans Orthop Res Soc, Atlanta, GA 1996.

- Equine emergency and intensive care: Case survey and assessment of needs (1992-1994). Proceedings, ACVS Annual Meeting, Chicago, 1995.

-Colonic luminal pressure in horses with strangulating or nonstrangulating obstruction of the large colon. Moore RM, Hance SR, Hardy J. ACVS Scientific presentation abstract. Vet Surg 23:411, 1994

- Mural blood flow distribution in the equine large colon during low-flow ischemia and reperfusion. Moore RM, Hardy J. Muir WW. ACVS Scientific presentation abstract. Vet Surg 23:411, 1994

-Effect of bacterial infection on equine synovial membrane cytokine production and cartilage metabolism. ACVS Scientific presentation abstract. Vet Surg 23:402, 1994

-Determination of mural blood flow distribution in equine large colon during low flow ischemia and reperfusion using colored microspheres. Moore RM, Hardy J. Muir WW. Fifth Equine Colic Research Symposium, Athens, GA 1994.

-In vitro effect of bacterial infection on synovial membrane hyaluronate synthesis and polydispersity, cytokine production and cartilage metabolism. Hardy J. Bertone AL, Malemud CJ. Transactions of the Orthopedic Research Society Annual Meeting, published in J Bone Joint Surgery 1995. -Physiologic responses of a novel isolated joint model. Bertone AL, Hardy J. Simmons EJ, Muir WW. Transactions of the Orthopedic Research Society Annual Meeting, published in J Bone Joint Surgery 1995.

-Pressure-volume relationships in the equine midcarpal joint. American Association of Biomechanics. Hardy J. Bertone AL, Muir WW. Jan 1995, Columbus, OH

-Synovial villi: distinct structural arrangement. Izumisawa Y, Tangkawattana P, Chang Y, Masty J, Hardy J. Bertone A, Yamaguchi M. American Society for Cell Biology 34th Annual Meeting, San Francisco, 1994.

-Effect of bacterial infection on hyaluronate synthesis by equine synovial membrane in vitro .Hardv J. Bertone AL^ Malemud CJ. Veterinary Orthopedic Society, Salt Lake City, Utah 1994.

-Effect of bacterial infection on cartilage metabolism in equine joints in vitro.Hardy J. Bertone AL^ Malemud CJ. Veterinary Orthopedic Society, Salt Lake City, Utah 1994.

-Determination of synovial membrane blood flow in equine joints using colored microspheres.Hardy J. Bertone AL, Muir WW. Am College Vet Surgeons, Veterinary Forum, San Francisco 1993.

-Care of the equine intensive care patientHardy J. Proceedings, Ohio Veterinary Medical Association annual meeting, 1993

-Diagnosis and management of colic in the foal.Hardy J. Proceedings, Congrès de l'A\%F, Paris France 1992.

-Fluid therapy in the management of colic in the horse. Hardy J. Proceedings, Congrès de I’AVEF, Paris France 1992.

-Arthrites septiques et ostéomyélites du poulain.Hardy J. Proceedings, Congrès de l'AVEF. Caen, France, 1991.

-Les boiteries de l'épaule. Diagnostic, pronostic et traitementHardy J. Proceedings, Congrès de Chirurgie Equine, Geneva, Switzerland, 1991.

-Effect of exercise on healing of articular cartilage in the horse. Hardy J. Bramlage LR, Weisbrode SE, Schneider RK, Bohanon TC, McKeever KM. American College of Veterinary Surgeons Ann Meeting, San Francisco, 1991.

-The significance of pre-training sesamoid radiographs on long term sooundness. Hardy J. Marcoux M, Breton L. American Association of Equine Practitioners annual meeting, Boston, Mass, 1989.

-Sesamoiditis in the Standardbred horse: a descriptive study.Hardy J. Marcoux M, Breton L.: Proceedings, XXJU World Veterinary Congress, Montreal, Canada. August 1987

-Medical treatment of colics.Hardy J. Proceedings of the Quebec Veterinary Association Annual Meeting, 1985.

IX FIELDS OF STUDY

Major field: Veterinary Clinical Sciences

Studies in Articular Physiology and Pathophysiology Alicia Louise Bertone, D.V.M., Ph.D., Adviser TABLE OF CONTENTS

Abstract...... ii

Acknowledgments ...... v

Vita ...... vi

List of figures ...... xiii

List of tables ...... xix

Chapters:

1. Introduction ...... 1

1.1 Nature of the pro b lem ...... 1 1.2 Hypotheses ...... 4 1.3 References ...... 5

2. Review of the literature ...... 8

2.1 Introduction ...... 8 2.2 General Anatomy and physiology ...... 8 2.3 Articular circulation ...... 11 2.4 Contribution of ischemia-reperfusion and generation of free radicals in arthritis...... 14 2.5 Role of nitric oxide ...... 15 2.6 Intraarticular pressure ...... 17 2.7 Sjmovial fluid dynamics ...... 20 2.8 Articular permeability and turnover of synovial fluid ...... 24 2.9 Functions of hyaluronan ...... 25 2.10 Role of articular innervation ...... 27 2.11 References ...... 34

3. Pressure-volume relationships in equine midcarpal joints ...... 39 3.1 Introduction ...... 39 3.2 Materials and methods ...... 41 3.3 Results ...... 46 3.4 Discussion ...... 48 3.5 References ...... 70

XI 4. Determination of synovial membrane blood flow in equine joints by colored microspheres and evaluation of the effect of increased intraarticular pressure .. 72

4.1 Introduction ...... 72 4.2 Material and methods ...... 74 4.3 Results ...... 81 4.4 Discussion ...... 83 4.5 References ...... 97

5. Local hemodynamics, permeability and O 2 metabolism of iimervated or denervated isolated joints in an il-1 model ...... 101

5.1 Introduction ...... 101 5.2 Materials and Methods ...... 102 5.3 Results ...... 112 5.4 Discussion ...... 115 5.5 References ...... 144

6. Cell trafficking, mediator release and articular cartilage metabolism in acute arthritis ofirmervated or denervated isolated jo in ts ...... 145

6.1 Introduction ...... 145 6.2 Materials and methods ...... 147 6.3 Results ...... 156 6.4 Discussion ...... 159 6.5 References ...... 191

7. Summary ...... 195

Appendix A Dynairtic vascular and trans-synovial forces of the isolated joint ...... 197 A.l Introduction ...... 197 A.2 Materials and methods ...... 199 A.3 Results...... 204 A.4 Discussion ...... 207 A.5 References ...... 237

Bibliography ...... 241

xu LIST OF FIGURES

Figure Page

2.1 Illustration to summarize the pathophysiologic mechanisms involved in joint injury ...... 33

3.1 Illustration of forelimb of horse, demonstrating position of needles for . measurement of intra-articular pressure in midcarpal joints ...... 53

3.2 Pressure-volume relationships in equine midcarpal joints demonstrating sigmoid shape of curve and difference between sub- and supra-atmospheric pressures. Elastance is significantly lower (greater compliance) at 135° than 90° angle (P=0.0016) ...... 55

3.3 Pressure-volume relationships in equine midcarpal joints at subatospheric (normal )pressures. Elastance is significantly lower (greater compliance) at 135° than 90° angle (P=0.028) ...... 57

3.4 Elastance as function of lAP at joint angle of 135°. Regression slope is steeper below than above atmospheric pressure ...... 59

3.5 PV relationships in equine midcarpal joint during infusion or withdrawal of saline or synovial fluid at joint angles of 90° (A) or 135° (B). Hysteresis calculated as: (area under infusion curve-area under withdrawal curve)/area under infusion (in %), is significantly greater with synovial fluid than saline (P<0.005) and at 90° than at 135° angle (P<0.005) ...... 61

3.6 PV relationships in equine midcarpal joints at 135° infused with isotonic saline or synovial fluid. Elastance was significantly greater with saline than with synovial fluid (P=0.045) ...... 63

xm 3.7 A: PV relationships at 3 low-pressure cycles (<30 mmHg) followed by 3 high-pressure cycles (30-80 mmHg) and at 3 high-pressure followed by 3 low-pressure cycles. B: inset of A; shows that elastance significantly increased at high-pressure cycles after prior distention at low pressures ...... 65

3.8 Relaxation of lAP as a function of natural log of time for 20-mmHg increments of LAPS (joint angle 135°). Slope of lAP-relaxation (lAP-R) curve is significantly increased with increasing lAP (P< 0.005) ...... 67

3.9 LAP-R as a function of LAP. There is a significant increase in slope of LAP-R curve with increased initial LAP. lAP = -8.37 + 3.00 x In (lAP) (R2 = 0.99, P< 0.0001) ...... 69

4.1 Medial aspect of the left forelimb of a horse illustrating normal anatomy and instrumentation for microsphere determination. The scale indicates approximate height above the heart of the metacarpophalangeal joint. . 88

4.2 Synovial membrane blood flow in the dorsal pouch of normal equine midcarpal joints in dorsal recumbency at a joint angle of 135° after increased intraarticular pressure (LAP) (Group 1) or in normal joints (Group 2). Data are mean +/-SE. * indicates significant difference from baseline ...... 90

4.3 Scattergram of the coefficient of variability between three simultaneous blood flow determinations obtained with three different microsphere colors, plotted against number of microspheres. This graph illustrates the higher variability associated with low microsphere numbers ...... 92

4.4 Bland and Altman plot of the difference between three serial (one minute interval) blood flow determinations in the same tissue specimen, to illustrate the repeatability of blood flow determinations. Indicated are the mean difference and 95% confidence interval for serial blood flow determinations ...... 94

5.1. Illustration of the isolated pump-perfused auto-oxygenated isolated joint model used in the study ...... 121

XIV 5.2. Circuit arterial pressure (A) and total vascular resistance (B) among groups and across time in isolated metacarpophalangeal joints from Baseline to 60 minutes. (Group 1 - ■ Control; Group 2-# Control-Denervated; Group 3- ▲ Inflamed; Group 4- ♦ Inflamed-denervated). There was no significant difference among groups ...... 122

5.3 Total vascular resistance (TVR) of among groups and across time in isolated metacarpophalangeal joints. (Group 1 - ■ Control; Group 2-# Control-Denervated; Group 3- ▲ Inflamed; Group 4- ♦ Inflamed-denervated) There was a significant decrease in TVR over time in all groups (P=0.(X)9), and there was no significant difference detected among groups ...... 124

5.4 Synovial membrane blood flow determined by colored microspheres among groups and across time in isolated metacarpophalangetd joints. (Group I - ■ Control; Group 2-# Control-Denervated; Group 3- ▲ Inflamed; Group 4- ♦ Inflamed-denervated) *Synovial blood flow was significantly increased (P=0.013) at 60 and 330 minutes compared to baseline and there was no significant difference detected among groups ...... 126

5.5 Synovial fluid production among ^oups from 240 to 330 minutes in isolated metacarpophalangeal joints.* Indicates a significant difference between inflamed (groups 3 and 4) and control joints (Groups 1 and 2) (P=0.001) ...... 128

5.6 Oxygen delivery among groups and across time in isolated metacarpophalangeal joints. (Group 1 - ■ Control; Group 2-# Control-Denervated; Group 3InAamed; Group 4- ♦ Inflamed-denervated) DÜ2 significantly increased across time (P<0.(X)01). There was no significant difference detected among groups ...... 130

5.7 Oxygen consumption (VO 2) and extraction ratio (0% ER) among groups and across time in isolated metacarpophalangeal joints. (Group 1 - ■ Control; Group 2-# Control-Denervated; Group 3- ▲ Inflamed; Group 4- ♦ Inflamed-denervated) * Indicates a significant difference between control (Groups 1 and 2) and inflamed joints (Groups 3 and 4) for VO?) (P=O.Ol) and 02ER(P=0.04)...... "._____ 132

XV 5.8 Photomicrographs (20 x) of laser argon acquired images of evans-blue albumin fluorescence of synovial membrane among groups and at fibrous or villous sites, demonstrating depth of absorption. Mean depth of absorption in the villous region for inflamed groups (groups 2 and 4) was 964.85 ± 141.55 p.m ...... 133

5.9 FTTC-dextran concentration in circuit plasma effluent among groups and across time in isolated metacarpophalangeal joints from 330 to 360 minutes. (Group 1 - ■ Control; Group 2-# Control-Denervated; Group 3- ▲ Inflamed; Group 4- ♦ Inflamed-denervated) * Indicates a significant difference between control (Groups 1 and 2) and inflamed joints (Groups 3 and 4) (P=0.02) ...... 135

6.1 Illustration of the isolated pump-perfused auto-oxygenated isolated joint model used in the study ...... 166

6.2 Synovial fluid total nucleated cell count across time among groups. (Group 1 - ■ . . Control; Group 2-# Control-Denervated; Group 3- ▲ Inflamed; Group 4- ♦ . . . Inflamed-denervated) *indicates a significant difference between control joints (Groups 3 and 4) and inflamed joints (Groups 1 and 2) (P=0.001) ...... 167

6.3 Synovial fluid total nucleated cell differential (%) over time among groups. There was a significant increase in % neutrophils and a significant decrease in % large and small monoculear cells across time. There was no difierence detected among groups ...... 169

6.4 Synovial fluid interleukin IB concentration ratios across time and among groups. *indicates a significant difference compared to baseline (P=0.001). ¥ indicates a significant difference between groups 3 and 4 (P=0.02). 171

6.5 Synovial fluid interleukin 6 concentration ratio across time and among groups. *indicates a significant difference between control joints (Groups 1 and 2) and inflamed joints (Groups 3 and 4) (P=0.01). . . . 173

XVI 6.6 PGE2 ratio over baseline across time and among groups. (Group I - ■ Control; Group 2-# Control-Denervated; Group 3- ▲ Inflamed; Group 4- ♦ Inflamed-Denervated) * indicates a significant difference in Group 4 (Inflamed-Denervated) (P=0.038) ...... 175

6.7 Total substance P (pg) in synovial fluid collection from 240-3(X) minutes (■ ) and 300 to 330 minutes (■).* Indicates a significant difference between control joints ...... (Groups 1 and 2) and inflamed joints (groups 3 and 4) (P=0.025) ...... 177

6.8 Synovial membrane histology specimens (1(X)X) illustrating A-a control isolated joint from group 1 and B- an inflamed isolated joint from group 3. Inflamed isolated joints (Groups 3 and 4) showed a significant neutrophilic vasculitis (P=O.OI)...... 179

6.9 Synovial membrane substance P and nerve specific enolase immunostaining. There was no difference among groups as to the number of positively stained fibers...... 181

A. 1 Pump-perfused isolated metacarpophalangeal joint preparation ...... 218

A.2 Equine forelimb and metacarpophalangeal joint before joint isolation demonstrating vascular sites for microsphere studies ...... 220

A.3 Capillary pressure versus venous pressure during isogravimetric conditions. Capillary pressure (Pcap) was greater (p<0.05) at higher circuit venous pressure (Pvcir) [y = l-46x + 10.8; r = 0.87] ...... 222

A.4 Change in circuit arterial blood flow (Qadr) caused by manipulation of circuit arterial (Pacjr) and venous {Pvâi) pressure (p < 0.009). The first bar from the left (al49) is before joint isolation, the second bar is after joint isolation at isogravimetric conditions. Different letter superscripts are significantly different (p<0.05) ...... 224

xvii A.5 Synovial fluid production (drops/min) caused by manipulation of circuit arterial (Pacir) and venous (Pvdr) pressures (p<0.021). Different letter superscripts are significantly different ( p ^ .0 5 ) ...... 226

A.6 Preparation weight gain (g) caused by manipulation of circuit arterial (Pacir) and venous (Pvcir) pressures (p< O.CH Pacir: p < 0.57 Pvcir). Different letter superscripts are significantly different (p<0.05). Letters with a “t” . superscript tended to be different ( p c O .l) ...... 228

A.7 Tissue compliance (ml/mmHg) changes caused by arterial (Pa^r) (p<0.05) and venous Pvcir pressure manipulation. Different letter superscripts are significantly different (p<0.05) ...... 230

A.8 Vascular compliance (ml/mmHg) changes caused by arterial (Pacir) and venous (Pvcir) pressure manipulation (p«^.01). Different letter superscripts are . significantly different (p<0.05). ( = a trend ( pcO.l) to be different from Pv20. Letters marked with a “t” superscript tended to be different (p<0.1). . 232

X V lll LIST OF TABLES Table Page

4 .1 Blood flow to articular soft tissues of normal midcarpal and metacarpophalangeal joints in horses positioned in dorsal recumbency, with the midcarpal joint at a 135” angle. Data are mean +/-SE. Syn = synovial membrane (synovium and subsynovium). Fibrous = fibrous joint capsule. * Midcarpal joint fibrous capsule blood flow significantly lower than midcarpal joint synovial membrane (P =0.034). ** Dorsal metacarpophalangeal joint blood flow significantly lower than palmar (P =0.048) ...... 96

5.1 Systemic hemodynamics and blood gas analysis (mean ± SE) among groups in horses submitted to isolated joint preparation ...... 138

5.2 Local hemodynamics (mean ± SE) among groups and across time in isolated joints ...... 139

5.3 Synovial fluid total protein, pH, Pa02, PaC02, 0 2 gradient, permeability surface area product and intraarticular pressure among groups and across time in isolated jo in ts ...... 140

5.4 Synovial membrane wet/dry ratio among groups in isolated joints. . . . 141

5.5 Depth of Evans blue-albumin fluorescence (pm) among groups in isolated joints ...... 142

6.1 Score description for evaluation of synovial membrane histology ....183

6.2 Synovial fluid II16 concentration, content and ratio among groups and across time. Ratio calculated using Group 1 as reference ...... 184

6.3 Synovial fluid 116 concentration, content and ratio among groups and across time. Ratio calculated using Group 1 as reference ...... 185

6.4 Proteoglycan synthesis (CPM/|ig protein) and content (p.g/p.g protein) and respective ratio of isolated to contralateral control joint among groups and at 18 hours, 72 hours and 144 hours time in culture ...... 186

6.5 Cartilage total glycosaminoglycan, chondroitin and keratan sulfate concentrations among groups at culture time 18,72 and 144 hours. . . . 187

XIX 6.6 Newly synthesized proteoglycan released in 24,48 and 72 hr media among groups expressed as amount released (CPM), ratio of isolated to contralateral control joints, and % of total cartilage content ...... 188

6.7 Endogenous proteoglycan released in 24,48 and 72 hr media among groups expressed as amount released (pg), ratio of isolated to contralateral control joints, and % of total cartilage content ...... 189

6.8 Histopathology scores (mean ± SB) among groups, sites, and comparing contralateral control to isolated joints ...... 190

A. 1 Signalment of horses used in the isolated metacarpophalangeal joint preparation (n=7) ...... 234

A.2 Systemic hemodynamics and isolated joint vascular function (mean ± SEM) in 7 horses ...... 235

A.3 Isolated metacarpophalangeal joint vascular and fluid values (mean ± SEM) at isogravimetric conditions ...... 236

XX CHAPTER 1

INTRODUCTION

Nature of the problem

Lameness is the leading cause for decreased performance, days lost to training and

wastage in h o rse s. i-2 Lameness attributable to joint disease represents greater than 50% of

lameness diagnosis.i.3 Ten years ago, it was estimated that the cost to the racehorse

industry alone was greater than half a billion dollars.^

In people, osteoarthritis is the most common form of joint disease. Its prevalence is

greater than 75% in people over 70 years old, and radiographic evidence of OA can be

found in as inany as 80% of people over 55 .5,6 Joint disease is the leading cause of

morbidity in people, particularly the elderly. Joint disease, initiated by trauma, is also the determinant of retirement from athletic endeavors.?

Joint disease has been classified by several schemes. A distinction has been made

between arthritis, where an inflammatory component predominates, and arthrosis in which

degeneration without evidence of inflammation is observed. On an anatomical basis, joint

disease has been classified based on the pathologic tissue type, as in synovitis,

osteochondral fracture, or cartilage degeneration. Duration of disease can qualify the

syndrome as acute, subacute or chronic, and stable or progressive. An etiologic basis for

classification of joint disease is most commonly used, because the disease process is complex, and may involve multiple tissues within the joints, and affect multiple systems(for example lupus erythematosus) 8 Joint disease has been classified as primary or secondary. Primary joint degeneration is a recognized syndrome in people, where joint degeneration is present without a specific etiology. Primary joint degeneration is not a recognized entity in horses, where joint disease can be traced to a specific etiology.

Secondary joint disease follow such etiologies as trauma, infection, immune-mediated related problems. Traumatic arthritis has been further specified as traumatic synovitis and capsulitis (type 1 traumatic arthritis), sprains and luxations (type HA traumatic arthritis), meniscal tears (type HE traumatic arthritis) or intraarticular firactures (type HC traumatic arthritis).8 Immune-mediated joint disease is rarely encountered in the horse, whereas it is a multi-factorial complex commonly recognized in people. All forms of arthritis, given enough time, can lead to chronic degenerative joint disease, also referred to as osteoarthritis or osteoarthrosis.

Models of joint disease have been defined and used in several animal species.

Models of osteoarthritis,9.10 infectious arthritis, n-i3 traumatic arthritis, 14 synovitis, and rheumatoid arthritis abound, 15 and even genetically engineered mouse models, predisposed to developing joint degeneration, exist. I6 These models aim to mimic a naturally occurring condition of people, but extrapolation or external validation of these studies and their conclusion should be taken with caution because of such issues as animal species, age, size, weight, gait, posture, and health, i? For example, rabbits have been shown to heal cartilage defects with hyalin cartilage, 14 whereas in horses and people, fibrocartilaginous tissue predominates in the healing defect. I7.i 8 Furthermore, this tissue is unable to withstand the rigors of exercise, and one year follow-up studies have revealed degeneration of the filling tissue. 19 Young animals have significantly greater ability to heal cartilage, because of greater endogenous activity of chondrocytes, than older animals. 20.21 Active motion, and to a greater degree, passive motion exercise have been shown to greatly increase the healing of cartilage defects. Finally, animals with endocrine problems such as diabetes, have decreased wound healing, and more relevantly decreased cartilage healing.23

Because of the importance of joint disease in the horse industry, many equine models of joint disease have been d esc rib ed . 17,18.24-28 These models aim to mimic synovitis, look at healing of cartilage defects, mimic naturally occurring osteochondral fractures, infectious arthritis, and most commonly are aimed at investigating therapies.

Recently, in vitro models have been explored, in order to provide controlled experimental conditions and maximize use of a n im a ls. 20,2 1,29-32 Extrapolation to the in vivo situation is more remote in these latter models, but a greater understanding of disease processes and pathophysiology can be gained at minimal cost and morbidity.

Many models of joint disease address a specific question, and examine a specific tissue or phenomena within the joint. Studies usually focus on the tissue rather than the entire joint as an organ. In reviewing the physiology literature, many aspects of joint physiology such as blood flow, transsynovial fluid exchange, and innervation have not been examined in the horse, and have been only partially described in other species.33-40 in other organ system models, such as heart, lung, liver and intestine it has become evident that many physiologic mechanisms and tissues interact in disease. Such an approach however, had not been examined in the joint. In other organ systems, ex vivo analyses, which allowed manipulation of physiologic mechanisms, were investigated, in order to improve our understanding of organ function, organ homeostasis, and disease process.

Classic models, such as the Krogh model,4i the Pappenheimer preparation,42 and the

Langendorff heart preparation,43 to name a few, were developed to study organ physiology and pathophysiology while controlling other systems. Our present study investigated the organ physiology of the joint.

H ypotheses

Our first hypothesis was that we could successfully develop an isolated joint organ preparation that would allow measurement and manipulation of the major Starling’s forces regulating capillary exchange, namely capillary, interstitial and articular hydrostatic pressures, and capillary, interstitial and joint oncotic pressure.'^! Our second hypothesis was that manipulation of Starling’s forces in the isolated joint would allow the calculation of the forces that regulate capillary fluid exchange to and from the joint cavity. Our third hypothesis was that interleukin-16 could be used as a model of acute inflammation in the isolated joint and that measurable alterations in articular metabolism, cell trafficking, cartilage chondrocyte metabolism and release of inflammatory mediators would result. Our fourth hypothesis was that denervation of the joint would modify the inflammatory response.

Our preliminary studies (Chapters 2 and 3) were designed to overcome and address several technical issues that pertain to an isolated joint organ preparation, such as the relevance of intraarticular pressure to hemodynamics, and the hemodynamic alterations associated with creating a surgical preparation. Appendix A describes the physiologic responses and perfusion of isolated joint preparation. Initially isolation of the equine carpus was attempted, with failure to produce a closed circuit. Extensive soft tissue trauma accompanied total carpal extracorporeal isolation, and partial isolation resulted in significant bone venous outflow and failure to isolate and control venous pressures. The equine metacarpophalangeal (MCP) joint was subsequently used successfully (Appendix A). In chapters 4 and 5 the isolated MCP joint organ was used in an experimentally induced acute arthritis model, to study the physiologic responses to inflammatory insult and the role of innervation in the acute response to this insult. R eferences

1. Jeffcott, L. B., P. D. Rossdale, J. Freestone, et al. An assessment of wastage in Thoroughbred racing from conception to 4 years of age. Equine VetJ 14:185-198, 1982.

2. Rossdale, P. D., R. Hopes, D. Wingfield, N J, et al. Epidemiological study of wastage among racehorses 1982 and 1983. VetRec 116:66-69, 1985.

3. Todhunter, R. J., G. Lust. Pathophysiology of synovitis: clinical signs and examination in horses. Compend Contin Educ Pract Vet 12:980-992, 1990.

4. Marvin, M. North american lameness survey. The Harness Horse 12-13,45, 1985.

5. Office, G. P. National Center for Health Statistics: Prevalence of osteoarthritis in adults by age, sex, race, and geographic area: United States 1960-1962. Vital and Health Statistics 15, Series 11:.

6. Brandt, K. D. Osteoarthritis: Clinical patterns and pathology. In: Kelley WN, Harris ED, Ruddy S, Sledge B, ed. Textbook of Rheumatology. Philadelphia; WB Saunders; 1985; 1432-1448.

7. Lane, N. E., J. A. Buckwalter. Exercise: A cause of osteoarthritis? Rheum Dis North Amer 19:617-633, 1993.

8. Mcll wraith, C. Chapter 7. Diseases of joints, tendons and related structures. In: Stashak TS, ed. Adam'j Lameness in Horses. Philadelphia; Lea & Febiger; 1987;.

9. Myers, S. L., K. D. Brandt, B. L. O'Conner, et al. Synovitis and osteoarthritic changes in canine articular cartilage after anterior cruciate Ugament resection. Arthritis Rheum 33:1406-1415, 1990.

10. Pelletier, J. P., J. M. Pelletier, L. G. Mnaymneh, et al. Role of synovial membrane inflammation in cartilage matrix breakdown in the Pond-Nuki model of osteoarthritis. Arthritis Rheum 28:1985.

11. Smith, R. L., D. J. Schurman. Bacterial arthritis. A Staphyloccoccal proteoglycan- releasing factor. Arthritis Rheum 29:1379-1385, 1986.

12. Smith, L. R., T. C. Merchant, D. J. Schurman. In vitro cartilage degradation by Escherichia coli and Staphylococcus aureus. Arthritis Rheum 25:441-446, 1982.

13. Riegels-Nielsen, P., N. Frimodt-Moler, J. Jensen. Rabbit model of septic arthritis. Acta Orthop Scand 58:14-19, 1987.

14. Furukawa, T., D. R. Eyre, S. Koide, et al. Biochemical studies on repair cartilage resurfacing experimental defets in the rabbit knee. J Bone Joint Surg 62-A:79-89, 1980. 15. Horn, J. T., A. M. Bendele, D. G. Carlson. In vivo administration with D-1 accelerates the development of collagen-induced arthritis in mice. J Immunol 141:834- 841, 1988.

16. Helminen, H. J. An inbred line of transgenic mice expressing an internally deleted gene for type n procollagen (COL 2A1). Proc Nat Acad Sci 92:582-595, 1993.

17. Desjardins, M. R. Cartilage healing: a review with emphasis on the equine model. Can VetJ 31:565-572, 1990.

18. French, D. A., S. M. L. Barber, D H, C. E. Doige. The effect of exercise on the healing of articular cartilage defects in the equine carpus. Vet Surg 18:312-321, 1989.

19. Howard, R. D., C. W. McDwraith, G. W. Trotter, et al. Long-term fate and effects of exercise on sternal cartilage autografts used for repair of large osteochondral defects in horses. Am J Vet Res 55:1158-1167, 1994.

20. Hjerquist, S. O., R. Lemperg. Histological, autoradiographic and microchemical studies of spontaneously healing osteochondral articular defects in adult rabbits. Calc^ Tissue Res 8:54-72, 1971.

21. Morris, E. A., B. V. Treadwell. Effect of interleukin 1 on articular cartilage from young and aged horses and comparison with metabolism of osteoarthritic cartilage. Am J Vet Res 55:138-146, 1994.

22. Salter, R. B., D. F. Simmonds, B. W. Malcolm, et al. The biological effect of continuous passive motion on the healing of full-thickness defects in articular cartilage. J Bone Joint Surg 62-A: 1232-1251, 1980.

23. Rosenbloom, A. L., J. H. Silvestein. Connective tissue and joint disease in diabetes mellitus. Endocrinology and metabolism clinics of North America 25:473-483, 1996.

24. Bertone, A. L., C. W. Mcll wraith, R. L. Jones, et al. Comparison of various treatments for experimentally induced equine infectious arthritis. Am J Vet Res 48:519- 529, 1987.

25. Gustafson, S. B., G. W. Trotter, R. W. Norrdin, et al. Evaluation of intra-articularly administered sodium monoiodoacetate-induced chemical injury to articular cartilage of horses. Am J Vet Res 53:1193-1202, 1992.

26. Hurtig, M. B. Use of autogenous cartilage particles to create a model of naturally ocurring degenerative joint disease in the horse. Equine VetJ Suppl 6:19-22, 1988.

27. Palmer, J. L., A. L. Bertone. Experimentally-induced synovitis as a model for acute synovitis in the horse. Equine Vet j 26:492-495, 1994.

28. Peloso, J. G., J. A. Stick, J. P. Caron, et al. Effects of hylan on amphotericin- induced carpal lameness in equids. Am J Vet Res 54:1527-1534, 1993.

29. May, S. A., R. E. Hooke, P. Lees. Adverse conditions in vitro stimulate chondrocytes to produce prostaglandin and stromelysin. Equine VetJ 23:380-382, 1991. 30. May, S. A., R. E. Hooke, P. Lees. Interleukin-1 stimulation of equine articular cells. Res Vet Sci 52:342-348, 1992.

31. McDonald, M. H., S. M. Stover, N. H. Willits, et al. Regulation of matrix metabolism in equine cartilage explant cultures by interleukin 1. Am J Vet Res 53:2278- 2285, 1992.

32. Morris, E. A., S. Wilcon, B. V. Treadwell. Inhibition of interleukin 1-mediated proteoglycan degradation in bovine articular cartilage explants by addition of sodium hyaluronate. Am J Vet Res 53:1977-1982, 1992.

33. Blake, D. R., J. Unsworth, J. M. Outhwaite, et al. Hypoxic-reperfusion injury in the inflamed human joint. Lancet 289-292, 1989.

34. Geborek, P., K. Forslind, F. A. Wollheim. Direct assessment of synovial blood flow and its relation to induced hydrostatic pressure changes. Ann Rheum Dis 48:281-286, 1989.

35. Knight, A. D., J. R. Levick. The influence of blood pressure on trans-synovial flow in the rabbit. J Physiol (Land) 349:27-42, 1984.

36. Knight, A. D., J. R. Levick, J. N. McDonald. Relation between trans-synovial flow and plasma osmotic pressure, with an estimation of the albumin reflection coefficient in the rabbit knee. Q J Exp Physiol 73:47-65,1988.

37. Levick, J. R. Fluid transport across synovial tissue in vivo [proceedings]. J Physiol (Lond) 1978.

38. Levick, J. R. Contributions of the lymphatic and micro vascular systems to fluid absorption from the synovial cavity of the rabbit knee. J Physiol (Lond) 306:445-61, 1980.

39. Levick, J. R. Joint pressure-volume studies: their importance, design and interpretation. J Rheumatol 10:353-7, 1983.

40. Levick, J. R. Synovial fluid and trans-synovial flow in stationary and moving normal joints. In: Helminen HI, Kiviranta I, Tammi M, Saamanen A-M, Paukkonene K, Jurvelin J, ed. Joint loading. Bristol; Wright; 1987; 148-186.

41. Taylor, A. E. Capillary fluid filtration. Starling forces and lymph flow. Circ Research 49:557-575, 1981.

42. Pappenheimer, J. R., A. Soto-Riviera. Effective osmotic pressure of the plasma proteins and other quantities associated with the capillary circulation in the hindlimbs of cats and dogs. Amer J Physiol 152:471-491, 1948.

43. Doring, H. J., H. Dehnert. The isolated perfused heart according to Langendorff. March, Germany; Biomesstechnik; 1979;. CHAPTER 2

REVIEW OF THE LITERATURE Introduction

Until recently, emphasis has been placed on articular cartilage biochemistry and cellular biology to gain understanding of the pathophysiology of joint disease and as a target for therapeutic efforts. In a recent paper, an extensive review of articular biochemistry in health and disease clearly summarized the importance of biochemistry and molecular biology to describe the pathophysiology of joint disease. i

The joint is an organ composed of several heterogenous tissues, with local blood supply, innervation, and fluid exchanges which function to maintain health and may contribute to the different aspects of disease. This paper will describe the normal articular physiology including blood flow, innervation, fluid exchanges between compartments and situate these physiologic processes when altered by disease.

General Anatomy and physiology

Tissues of the joint cavity have a specialized composition and three dimensional anatomy which ultimately relate them to their specific biomechanical function. As such the articular cartilage, whose ultimate function is to absorb and transfer loads, is a composite of collagen which grants it its tensile strength and highly charged proteoglycan subunits which provide compressive stifftiess. In order to maintain cartilage composition and function, a slow but steady turnover of its components occurs, the rate of which is dependent onspecific factors including age, mechanical load, and joint environment.2-5 The building

8 materials necessary for this turnover are provided through blood flow and exchanges fromthe synovial membrane and joint capsule, as well as through ultrafiltration and formation of synovial fluid, which provides the ultimate media for these exchanges to occur.6

The supply of nutrients to the avascular but metabolically active articular cartilage is provided by exchanges through the synovial microcirculation and transport in synovial fluid. The synovial membrane is a specialized unit designed to provide a pathway fo fluid exchanges, as well a blood supply, and add to the composition of synovial fluid through hyaluronate synthesis. The synovial membrane is composed of an intima (also referred to as synovium), consisting of luminal layer I to 5 cells deep, a capillary plexus which lies

6-11 pm beneath the intimai surface, and a deeper network of lymph vessels, and a subintima (or subsynovium) composed of adipose, areolar or fibrous tissue.6 The intima is designed to favor exchanges between capillaries and the joint cavity: it lacks a basement membrane, intercellular gaps are present (up to 19% in the rabbit knee), and intimai capillaries are fenestrated, with fenestrae are oriented towards the joint cavity .6

Furthermore, the synovial intima adopts a three dimensional architecture which is dependent on its biomechanical environment. In areas of high biomechanical stress, the synovium is flat and rests on a fibrous mechanically strong subintima. This is particularly true of synovium underlying tendons that cross over a joint, such as the extensor tendon overlying the metacarpo-phalangeal joint. In synovial recesses, the intima is thrown into a three dimensional villous architecture. This network is more richly vascular and has been shown to be a preferential site of solute and macromolecule exchanges.? Hyaluronan synthesis is also greater in synovium from synovial villi.8 Intimai cells include synovial fibroblasts (or type B cells, approximately two thirds of cells) and bone marrow derived mononuclear phagocytes (or type A cells, approximately one third of cells)9. Light microscopy techniques are not a useful guide for assessing lineage, as subintimal macrophages are often elongated, and intimai fibroblasts may take a rounded appearance.9

These cells can be separated on the bases of cytochemical staining for uridine diphosphoglucose dehydrogenase, a marker for hyaluronan synthesis, and non-specific esterase, a macrophage marker, lo These cells are also unique as they are found in close association with specialized components of the intimai matrix. As an example, the adhesion molecule ICAM-1, expressed by synovial fibroblasts, binds to hyaluronan. Although technical difficulty has made the identification and relative numbers of intimai cells of different lineage, there is currently no evidence to suggest that intimai fibroblast can transform to a macrophage phenotype.9 Although the synovial intima is interrupted and does not possess a basement membrane, interaction of synovial fibroblasts with the specialized matrix of the synovial intima forms a barrier adapted to the regulation of synovial fluid composition. Because of its high hyaluronan content, the synovial intima forms a continuum with synovial fluid for exchanges of fluids and solutes.^ In addition, other glycoproteins such as fibronectin, laminin, type IV collagen, type V collagen, entactin, and sulfated glycoaminoglycans such as chondroitin sulfate, secreted by the deeper type B synoviocytes, may serve to anchor the intimai cell layer to the underlying connective tissue. ^ The ability of the intima to maintain itself as a layer appears closely related to cell- matrix-cell interactions, through expression of adhesion molecules such as

ICAM-1, VCAM-1, fibronectin, laminin and others and their integrin ligands are likely candidates. With inflammation, the relative predominance of these cells types is altered, with subintimal macrophages forming 50-70% of cells in the intima. Integrin expression is also increased, and the morphology of the cells is changed from small cells arranged parallel to the joint surface, to a more superficial location arranged perpendicular to the joint cavity. These cells are also the dominant source of intimai vascular cell adhesion molecule

10 (VAM-1)9 Expression of cell adhesion molecules by synovial intimai cells serve to direct leukocyte trafficking in disease. Most multinucleate cells within inflamed synovial intima carry macrophage lineage markers.9 There is also evidence that synovial intimai cells could direct leucocyte trafficking in health and disease, through adhesion molecule expression.9

Currently, there is no evidence that subintimal macrophages can divide. Intimai fibroblasts do divide in disease, albeit at a slow rate, consistent with synovial proliferation observed with chronic inflammation. 12

Articular circulation

The synovial membrane functions to maintain articular homeostasis by providing a pathway for the exchange of nutrients and metabolic by-products between blood and synovial tissues, including articular cartilage 13.14 Optimal oxygen delivery to articular tissues serves to maintain normal synovial fluid composition 15,16, chondrocyte metabolism, and ensure normal matrix composition and turnover 17-19. The efficiency of exchange between the synovial membrane capillaries and joint cavity is dependent on capillary density, capillary depth and blood flow [Levick, 1990 #41].

Several tissues in the joint are provided with a rich vascular supply which can be affected by physiologic or pathologic states. These include the joint capsule, synovial membrane, intra and periarticular ligaments, and subchondral bone. Articular cartilage is avascular, but the outer third of the menisci is supplied by small vessels arising from the joint capsule. The articular blood supply of most diarthrodial joints is formed by small branches of the epiphyseal arteries running at the junction of the periosteum and the synovial membrane, forming an arterial circle (circulus arteriosus). Larger branches penetrate the bone whereas smaller branches remain at the periphery of the articular cartilage, forming the perichondral circulation. Subchondral bone blood supply is provided

11 by the epiphyseal arteries, which travel in the epiphysis parallel to the articular cartilage, giving off at interval small perpendicular branches which end in capillary loops at the deep surface of the calcified cartilageÆefore physeal closure, this epiphyseal circulation is distinct from the metaphyseal circulation.This segmentation of blood supply may explain the subchondral and epiphyseal location of infection in the equine n e o n a te . 20,21

Conversely, segregation of blood supply to the metaphysis in the older foal provides an explanation for localization of infection to the more mature side of the growth p late. 20,21

The synovial membrane vascular supply is composed of capillaries which are very sparse in areas of high mechanical stress. The angle of reflection of the synovial membrane is composed of a rich vascular plexus and synovial villi are incompletely penetrated by a central arteriole. The richest capillary density in the synovial membrane is within 2 5 pm of the joint surface. Capillary density is greatest in areolar or adipose synovium and lowest in fibrous synovium. Similarly, blood flow to synovium is greater than in the fibrous joint capsule .22 Synovial capillaries are fenestrated, with the fenestration oriented towards the joint cavity.6 The rich and superficial blood supply to the synovial membrane explains why hemarthrosis after needle puncture is frequently encountered, and provides an explanation for the hematogenous spread of infection to that area. In addition, the close proximity of highly vascular (synovium) to avascular (cartilage) tissue, referred to as a “borderline area”, makes this anatomical site a preferential location for systemicaUy injected particles.23 Many joint diseases exhibit greater inflammation at the angle of reflection of the synovium on the articular cartilage, and this site may be more appropriate when synovial membrane biopsies are obtained for culture in septic arthritis.24 In addition, villous synovial membrane is more vascular than fibrous, and preferential exchanges of small molecules such as albumin, as well as hyaluronan production, occurs in synovial villi.7.25

There is evidence that the synovial membrane is capable of handling a certain amount of

12 bacteria without developing infection.24^6 The superficial capillaries and small venules have thin fenestrated walls, akin to that observed in the kidney, intestine and choroid plexus, that are well suited for fluid transit.27 Deeper vessels have thick endothelium and are the site of action of histamine, and transvascular migration of leukocytes. Even in normal joints, deep venules have been shown to have high permeability to particles of up to

250 A , and the preferential localization of intravenously injected bacteria to the joint may be explained in part by this high permeability.27

Factors that can acutely affect synovial blood flow include intra-articular pressure

(lAP), local temperature, joint motion, vasomotor tone and reflexes, and local release of vasoactive mediators. Although large arteries supplying the joint have pressure approaching systemic arterial pressure, arterioles and capillaries of the synovial membrane have much lower pressure such that a relatively mild increase in lAP can cause significant tamponade.

In one study, a significant decrease in blood flow was measured when lAP reached 20 mm

H g.28 In the horse, a significant decrease in blood flow to the synovial membrane was measured after an increase lAP of 30 mm H g .22 In one study, there was an inverse relationship between intraarticular PO 2 and synovial fluid volumes with intraarticular acidosis developing at IAP<45 m m H g.29

Chronic pathologic processes can significantly decrease blood flow to the synovial membrane by two mechanisms: the increased fibrosis of the joint capsule can result in a significant decrease in capillary density; and this fibrosis can decrease compliance of the joint capsule, making it more susceptible to be affected by effusion.

The consequences of decreased articular blood flow include chronic ischemia, generation of lactate, synovial fluid acidosis, and generation of free radicals. Chronically hypoxic joints are more affected by exercise than normal joints. Finally, a decreased blood

13 supply can decrease drug delivery to the joint, and decrease clearance of metabolites from the joint.

Morphometric and morphologic analysis of synovial vessels provide an insight to the contribution of synovial blood flow in disease. Examination of normal synovial vessel morphology demonstrate capillary flattening with intraarticular pressure increases greater than 25 mm Hg, an indication that decreased blood flow, decreased clearance and decreased lymph flow can result from increased intraarticular pressure, and contribute to ischemia, edema, and joint effusion. In chronic arthritis decreased capillary density is observed, and increased intercapillary distance suggests altered transsynovial fluid exchanges, as well as relative ischemia. In immune-mediated conditions such as rheumatoid arthritis, vascular endothelial morphology directly correlated with disease status, with joints in active inflammatory stages showing changes characteristic of high endothelial venules, thus correlating well with the increased lymphocytic infiltration associated with these inflammatory stages.30

Contribution of ischemia-reperfusion and generation of free radicals in arthritis.

Generation of free radicals and subsequent membrane and tissue damage has been implicated in the pathophysiology of several disease processes including intestinal strangulation, spinal cord injury, myocardial injury, neoplasia and acute and chronic inflammation. In an attempt to explain the ongoing tissue damage observed in several forms of arthritis, a role of ischemia-reperfusion injury was proposed 3i and subsequently substantiated 32,33. Hypoxic-reperfusion injury is mediated predominantly by oxidative damage precipitated by reactive oxygen species, particularly the superoxide radical and its dismutation product hydrogen peroxide and hydroxyl radical. Generation of free radicals

14 originates in the generation of xanthine oxidase during the ischemic cycle, with free radical production during reperfiision. Xanthine oxidase has been immunolocalized in synovial endothelium,34 and production of free radical species by rheumatoid synovium has been documented.35 Free radicals can degrade articular cartilage matrix, hyaluronan, and cause chondrocyte death. Generation of free radicals is also a feature of the respiratory burst of polymorphonuclear leukocytes, which produce reactive oxygen species responsible for killing microbial pathogens. Thus cellular infiltration during inflammation can be detrimental through production of free radicals. Hypoxia within the diarthrodial joint may result from decreased arterial 0%, decreased synovial membrane permeability, decreased synovial blood flow, and/or increased 0% utilization through increased metabolic demand.29 In one study, oxygen consumption was significantly increased in joints with experimentally induced articular inflammation, and a concurrent increase in 0% extraction ratio was measured until delivery matched demand.? Cycles of ischemia-reperfusion can also be generated by joint effusion and increased intraarticular pressure, with subsequent decreased blood flow .22 The alteration in blood flow caused by effusion is worsened with exercise.33 Evidence for such mechanisms exist. Oxygen partial pressure in inflamed joint is higher than non-inflamed joints. In one study, oxygen extraction ratio and oxygen consumption was increased in equine joints inflamed with interleukin-1 (D-1) suggesting inability of delivery to meet demand.? These studies support rest and resolution of effusion as part of the treatment of joint disease.

Role of nitric oxide

Nitric oxide (NO) is a small molecule that is synthesized from the guanido group of the amino acid L-arginine by a variety of cells. A family of enzymes called the nitric oxide synthases (NOS) are involved in this reaction and two isoforms of the enzymes have been

15 identified: a constitutive Ca-dependent enzyme and an inducible Ca-independent enzyme.

Initially identified as endothelium-derived relaxing factor, NO was shown to have marked effect on vascular smooth muscle, causing vasodilation, and on platelets, neutrophils and macrophages to influence platelet aggregation, bacterial killing, and neutrophil trafficking.

Macrophages, peripheral blood monocytes, neutrophils and lymphocytes produce NO by means of the inducible form of NOS during host defense, inununological reactions and septic shock. This inducible form can be activated by endotoxin, or by inflammatory mediators such as interleukin-1, tumor necrosis factor and interferon (IFN), and can be inhibited by dexamethasone. Evidence of a role of NO in arthritis is increasing. Nitrite, the stable measurable end product of NO, has been measured in patients with rheumatoid arthritis and osteoarthritis. NOS inhibitors reduced the severity of several experimentally induced models of arthritis. In vitro studies have recently demonstrated that articular chondrocytes can release NO after stimulation with endotoxin and inflammatory mediators, and that this release could be inhibited by L-NAME, high concentrations of methotrexate and high concentrations of dexamethasone. Low levels of constitutive NOS activity were measured in synovial tissue, probably of endothelial origin, but further release could not be induced from that tissue under stimulation. Nitric oxide is highly toxic, but even more so when combined with superoxide free radical to produce the peroxynitrite anion, causing further tissue injury.36 In joints, NO decreases proteoglycan synthesis, and increases metalloprotease mediated collagen and proteoglycan degradation. NO also increases PGE 2 synthesis through direct activation of cyclo-oxygenase. The therapeutic benefit of NOS inhibition in arthritis stems from the fact that this pathway is upstream from the point of

NS AID anti-inflammatory effects of non steroidal drugs. Implication of NO in arthritis has been suggested, through involvement in the immune response, tissue injury and inflammation. In addition, NO is thought to be responsible for the vasodilator effects of substance P, and thus indirectly could play a role in plasma extravasation and joint

16 effusion. Manipulation of NO production may have some therapeutic benefits.

Glucocorticoids inhibit the induction of NO, and NSAID may inhibit production of NO by synovial fibroblasts. The ubiquitous nature of NO makes therapeutic targeting difficult and non-specific at this time. Identification and purification of isoforms that are specific to the diarthrodial joint may improve our ability to target NO in joint disease. 37

Intraarticular pressure

Intraarticular pressure (lAP) is below atmospheric pressure in most joints at the angle of ease and pressures between -2 to -12 cm H20 have been reported. Recordings of lAP in equine joints yielded similar subatmospheric values of for the midcarpal and metacarpophalangeal joints.38.39 Maintenance of this negative pressure is thought to occur by joint motion, which enhances lymph flow from the interstitium, and by joint flexion which promotes fluid absorption by raising lAP. The normal fluid balance is therefore maintained through 2 pumps in series, one that enhances fluid exchange to the interstitium, and one that enhances lymph flow. Examination of pressure-volume curves in normal joints indicates that this relationship is sigmoid, with low articular compliance at normal subatmospheric pressures, an increased compliance at supraatmospheric pressures up to 30 itimHg, and another increase in compliance at high IAP.39 This relationship can be explained as a resistance to joint distention at lAP in the normal range, followed by accommodation of effusion at lAP up to 30 mmHg. This may prevent collapse of synovium capillaries and preserve joint blood flow. At lAP > 30 mmHg, the decreased compliance may counteract further effusion and articular fluid accumulation. Rupture of the midcarpal joint capsule was documented at lAP of > 80 mmHg in horses, and this rupture was located in the palmar lateral pouch of the midcarpal joint, as has been observed clinically.

17 The determinants of lAP include joint capsule compliance, joint angle, previous distention history, compliance of the joint capsule, muscle tension, and joint load.40.4i in addition, determination of pressure-volume relationships are also dependent on the type of infiisate used for measurement.39.42 Chronically inflamed joints have thickening and fibrosis of the joint capsule, resulting in decreased compliance. Joint flexion increases lAP. Slow distention-compression cycles result in progressive stress-relaxation of the joint capsule and a gradual increase in articular compliance. However, if rapid successive infusion-withdrawal cycles are performed, a progressive decrease in articular compliance can be measured. If the joint is distended with synovial fluid, an increased compliance is observed compared to infusion with saline, probably because of the lack of a fluid interface which increases surface tension and promotes joint collapse.

Intraarticular pressure is an important determinant of synovial membrane blood flow and transsynovial fluid movement. Synovial membrane capillaries, which have a high density at a depth of approximately 25 pm, are affected by increased lAP. Studies have demonstrated a significant decrease in synovium blood flow at IAP> 20 m (rabbit) or 30 mm Hg

(horse)22,28. Decreased lA blood flow can lead to synovial fluid hypoxia, acidosis, and lactate accumulation.29 Chondrocyte metabolism is significantly altered by hypoxia.

Furthermore, increased lA pressure and exercise may lead to cycles of ischemia and reperfursion of the synovial membrane, with evidence of reperfusion injury. Superoxide generation, and other oxygen-derived metabolite have been demonstrated in synovial fluid after reperfusion in the synovial membrane. In mm superoxide can also affect chondrocyte metabolism.

Transsynovial fluid absorption is significantly affected by lAP. In the rabbit, transsynovial fluid movement increases 6-fold above a critical pressure of 9.5 cm H20 also termed the breaking pressure. These findings contradict Starling’s hypothesis, which predicts linear

18 relationship between fluid exchange and extravascular pressure.'*^ The clinical relevance of these findings are several fold. First, since flexion increase lAP, an increased lAP is expected with joint effusion and flexion, thus explaining the pain noted clinically with flexion tests performed on effused joints. Secondly, since cycles of reperfiision can be provoked with exercise in the presence of joint effusion, and that reperfiision and generation of free radicals can cause tissue injury, rest is indicated in joints with effusion.

The effect of history-dependance on lAP and compliance explains the lack of correlation between the volume of effusion and pressure generated from the effusion. Long term slowly accumulating joint effusion will have relatively low lAP compared to fast-developing effusions. In addition, pain due to effusion is caused by periarticular tension receptors, stress relaxation may explain why rapidly forming effusions are more painful than slower forming ones.

Compartmentation of the joint is the functional separation of joint compartment at physiologic pressure, and has been demonstrated in the rabbit stifle and the equine metacarpophalangeal joint. Because compartmentation was not observed in the non-weight bearing equine midcarpal joint, this would suggest that this phenomena is operative either in small joints or in large weight bearing joints.

The breaking point phenomena is not observed in edematous synovium either in animals with congestive heart failure or in acutely inflamed synovium, further supporting the observation that this phenomena is due to enlargement of the synovium interstitial channels.

This phenomena is also present with exercise, such that increased synovium conductance is observed for many hours following exercise. Therefore physiological injections studies of joints should be aware of this phenomena, and therapeutic joint injections should not be made after exercise.

Chronicity of joint effusion affects the magnitude of intraarticular pressure measured at rest and during exercise. Patients with acute traumatic joint effusions had a lower mean resting

19 intraarticular pressure and generated a significantly lower intraarticular pressure at exercise than patients with chronic effusions 44. These findings may be explained by the lower compliance of chronically inflamed joints. In chronically inflamed joints, pressures generated at rest but more remarkably at exercise are higher than capillary pressure and may reach values that are higher than systolic pressures. That severe synovial damage does not occur suggests the presence of protective mechanisms to prevent total ischemia to the synovium 44. Normal exercise does not produce significant fluxes in intraarticular pressures , emphasizing the critical role of effusions on the generation of intraarticular pressure 44,45. Interestingly, one of the most important mechanisms for lack of generation of high intraarticular pressures in acute traumatic effusions is quadriceps inhibition. The presence of this reflex has not been documented in horses.

Synovial fluid dynamics

Understanding the pathophysiologic mechanisms behind synovial fluid turnover becomes important when one considers the common clinical finding of joint effusion in joint disease, the use of synovial fluid markers as indicators of disease state and progression, and the use of systemic or intraarticular drugs in the treatment of joint disease.

The histology, ultrastructure and embryology of the synovium support the theory that the joint cavity is a modified interstitial space.46 The interstitium is composed of small proteoglycans, hyaluronan and fluid which is an ultrafiltrate of plasma. Similarly, joint fluid is an ultrafiltrate of plasma to which is added hyaluronan and is considered a form of interstitial fluid where 1) the concentration of small solutes is equal to plasma, after allowing for equilibrium for local metabolism and the Gibbs-Donnan effect; 2) the concentration of large solutes (colloids) is lower than that of plasma; 3) the rate of formation and absorption is based on Starlings hypothesis of fluid exchanges.4?

2 0 Current research supports a Starling based model for synovial fluid dynamics, which consists of ultrafiltration of plasma, passage of the ultrafiltrate to and from the synovial cavity via the synovial intima, and absorption into blood vessels and lymphatics.

The Starlings forces regulating fluid exchanges can be described by: J = K [(Pc-PO - C

(TCp-TCj)] where J=volume flow across the capillary, K is the filtration coefficient across the capillary wall (and is a function of capillary permeability (P) and surface area available for exchanges), Pc is the capillary hydrostatic pressure, P; is the interstitial hydrostatic pressure, ct is the reflection coefficient, Ttc is the capillary oncotic pressure, and tq is the interstitium oncotic pressure.48 This equation represents equilibrium at steady state. The dynamics governing fluid exchanges in the joint is best explained by a three-compartment model consisting of the vascular space, the interstitium and the joint space, which each membrane interface regulated by the sum of individual Starlings forces. Several studies have calculated the magnitude of individual forces across the joint and have identified several features of fluid dynamics that are unique to the joint. Increases in arterial and to a lesser degree, venous vascular pressures produces an increase in transsynovial fluid flow rather than interstitial flow. This can be explained by two mechanisms. The low reflection coefficient of the synovial membrane, which favors translocation of colloids into the joint, thus increasing tq, favors fluid flow into the joint. In addition, calculation of the transitional microvascular pressure which is the hydrostatic capillary pressure necessary to initiate synovial fluid or lymph flow yields a lower value for synovial fluid, thus again indicating preferential flow into the joint. The joint cavity therefore represents an overflow system in series with the vascular and interstitial space which can be opened at relatively low pressures. An example of this interaction serves to explain the relationship between the vascular interstitial and articular compartment. In immobilized joints, lymph flow is absent, such that fluid absorption from the joint through is preferentially through

21 capillaries. It can be demonstrated that in immobilized joint, there is steady increase in joint fluid volume, resulting in a mild increase in joint pressure, illustrating a preferential pathway of filtration to the joint space.

Synovial fluid colloids are an important driving force in joint fluid dynamics.

Articular albumin and hyaluronan concentrations are molecules that play a role in oncotic pressures.49 The coiled three dimensional hyaluronan network creates channels or excluded volumes that are though to be the mainstay of fluid exchanges within the joint. Because of the low synovial membrane coefficient, increases in intraarticular protein concentration are commonly observed with articular inflammation that leads to increased capillary permeability. In turn, this increases the oncotic intraarticular pressure participates in creating joint effusion. Hyaluronan concentration, but not molecular weight, has little influence on clearance of albumin from the joint, suggesting that other mechanisms, such as increased capillary permeability, and increased intraarticular pressure play a more significant role in joint clearance.50 The half life of the HA molecule in joints is relatively short (12-20 hours) suggesting that alteration in composition and molecular structure can quickly impact joint rheology. The dynamic state of synovial fluid composition raises questions as to the usefulness and pertinence of a one time measurement of markers of disease.

Increased synovial fluid protein concentrations results in two phenomena: increased oncotic pressure, and increased synovial fluid viscosity. Increased oncotic pressure decreases fluid absorption as could be predicted from Starling’s equation, and increased viscosity reduces flow through transintimai and leak channels. Hyaluronate also raises normal synovial fluid oncotic pressure in a ratio that is not proportional to its concentration.

For example, a 1 % hyaluronate solution doubles osmotic contribution firom protein. In joint effusions, oncotic pressure is raised, because of the increased total protein, but because

2 2 hyaluronate concentrations are low, total oncotic pressure is doser to the theoretical pressure due to protein.

Absorption of fluid from the joint.

The principal determinant of fluid absorption form the joint is intraarticular hydrostatic pressure and the sensitivity of flow (measured by absorption of fluid from the joint) to pressure increases almost six-fold above an intraarticular pressure of approximately 9.5 cm

H20, a limit which has been termed the breaking pressure or breaking point phenomena.5i

The experimental observation of a breaking point phenomena further yielded the theory that synovial conductivity was a constant below this limit, and a linear function of intraarticular pressure above breaking point. The clinical significance of this observation is that above a critical lAP, synovial membrane conductivity increases linearly with lAP to prevent the formation of effusion. Furthermore, interruption of lymphatic drainage does not affect this relationship.51 Lymphatic flow in the immobile joint is not a significant pathway for synovial fluid absorption. Turnover of synovial fluid is an important issue as synovial fluid concentration of a variety of macromolecules has been used to assess joint health in a variety of clinical conditions. In turn, concentration of these markers is determined by their rate of release from the tissue involved but also on factors that affect clearance of the molecule. These factors include volume of synovial fluid, which relates to intraarticular pressure and therefore hydraulic conductance, synovial blood flow, lymph flow, inflammation, joint motion and composition of synovial fluid.50.52 Fluid absorption from the joint occurs by two parallel pathways, the capillary bed and the interstitial spaces.

Bidirectional flow of fluid across the synovial membrane has been demonstrated, showing that fluid exchanges do occur at static fluid volumes, without the need for cyclical hydrostatic force changes dependent on muscle contraction.^

23 Articular permeability and turnover of synovial fluid

The ease of collection of synovial fluid in the horse has enabled the measurement of many molecules, including markers o f osteoarthritis, and therapeutic agents. In humans with OA, several molecules have been measured as markers of disease process, progression and response to treatment. These include keratan sulfate, chondroitin sulfate, proteoglycan core protein, type H collagen polypeptide, link protein, COMP, and others.

Serum concentration of some of these molecules, as an estimate of synovial fluid clearance and therefore concentration have also been used as markers. However, the concentration of a molecule in SF is related to its rate of release from cartilage, the volume of synovial fluid, and lymphatic or venous clearance from the joint. For example, one study detected a higher concentration of the 7-D-4 CS epitope and a significantly lower 5-D-4 KS epitope in knee joints with acute traumatic injuries. However, it is possible that the smaller KS epitope was cleared faster from the joint thus resulting in lower concentrations, rather than a reflection of altered cartilage synthesis or degradation.53 Even though hyaluronan molecules can form three-dimensional networks that can decrease the volume of distribution and clearance of small molecules such as albumin, synovial fluid hyaluronan concentration or molecular weight play little role in clearance of protein from synovial fluid.50 Low grade synovitis as seen in osteoarthritis can result in significantly increased clearance of small proteins such as albumin from the joint.54 Therefore the presence of synovitis is a significant variable which must be considered if response to therapy is measured by synovial fluid concentration of a marker protein.54 Recently, the preferential pathway of albumin exchange in equine fetlock joints has been localized to villous synovium, and a significant increase in albumin permeability was measured in experimentally induced articular inflammation. The same

24 study also identified a significant effect of denervation, with denervated joints showing a significant decrease in macromolecule (MW: 144,000) permeability? Lymphatic flow is thought to play an important role in protein clearance from the joint. Lymphatic flow, protein permeance and plasma protein clearance was greater in rheumatoid than osteoarthritic joints. Protein permeability was also inversely proportional to molecular radius, suggesting that proteins are cleared from the joint by diffusion. 55 Intraarticular local anesthetics, but not corticosteroids, increased the half life of Xe in rheumatoid knee joints, suggesting a local vasoconstrictive effect of the local anesthetic on synovial vascular tissue.56

Functions of hyaluronan

Hyaluronan is a non-sulfated glycosaminoglycans consisting of alternating units of

D-glucuronic acid and N-acetyl-D-glucosamine. It exhibits polydispersity, but the average molecular weight is in the order of several millions. In dilute solutions, each molecule behaves as a large coil, but as the concentration increases, entanglement of the coils occurs, eventually forming a uniform meshwork. Viscosity is non-linear and increases exponentially as the hyaluronan concentration increases, probably as a result of this three dimensional network. The effect of hyaluronate on synovial fluid viscosity is proportional to chain length, protein concentration, pH, ionic composition and temperature. Viscosity decreases rapidly with increased shear rate. Viscosity is variable between joints, being high in small joints. In a given joint viscosity varies inversely with volume. The viscoelastic nature and shear dépendance of hyaluronan solutions have been show, and a role in boundary lubrication assigned to the hyaluronan molecule. It is apparent however, that the nature and configuration of the molecule, and the existence of receptors to hyaluronan, provide some other insights as to the role of hyaluronan in vivo. Hyaluronan solutions

25 provide a barrier against water flow, and may act as a barrier against rapid tissue weight changes. The meshwork may also act as a sieve to regulate transport of macromolecules, and exclude macromolecules from space in the system. One theory proposed that the meshwork configuration and the rapid clearance of HA from the joint (20 hours in normal joints) may provide a scavenging function to this molecule. Finally the discovery of hyaluronan binding proteins, also referred to as hyaladherins, provided new insight as to the function of hyaluronan. The first and most known hyaladherin is aggrecans and link protein, which in combination with hyaluronan form the large aggregates of articular cartilage. Other surface receptor proteins, such as CD44 and receptor for hyaluronan mediating motility (RHAMM) may play a role in cell biology. Hyaluronan surface cell binding has been proposed as a mechanism to prevent cell fusion, and these receptors are the first to be recognized in the embryo, before joint cavitation, suggesting a role of hyaluronan to initiate joint formation. RHAMM is thought to activate an intracellular tyrosine kinase and initiate cell locomotion. Finally, cell endocytosis is also thought to be regulated by another hyaladherin, the lEC receptor. The half life of hyaluronan in joints in very low, and is similar for high or low molecular weight preparations. This half life decreases markedly with inflammation, but to date hyaluronan degradation has not been identified in the joint cavity. The liver appears to be a major site of hyaluronan degradation, with some occurring in periarticular tissues.^? Recently, synovial fluid hyaluronan has also been shown to play an important role in the mechanical properties of the joint cavity, such that the joint capsule is more compliant with synovial fluid than with salin e.39

A role of hyaluronan to decrease joint pain has been demonstrated in a bradykinin induced model of articular pain. In that model, hyaluronan with a molecular weight of greater than

40 kd was shown to have an analgesic effect, and hyaluronan of 860 kd and 2300 kd

26 produced high and long acting analgesia (72 hours after injection). This effect was not related to binding to hyaluronan receptors or bradykinin receptors.58

There is good experimental and theoretical evidence to support a role for hyaluronan to inhibit rapid efflux of fluid from synovial cavities at increased intra-articular pressures, by its impaction in the synovial intimai matrix.9

Finally, hyaluronan binding protein CD44 and the receptor for hyaluronan mediating motility (RHAMM) suggests an important role of hyaluronan in intercellular reactions and in the immune response and cell traffic.^

Role of articular innervation

The joint is an organ that is richly supplied with large myelinated afferent and efferent nerve endings and small unmyelinated C-fibers. Sensory and motor innervation maintain joint stability through reflex afferent, such that in the absence of these protective reflexes, severe arthropathy may develop if the joint is made unstable.59 Loss of sensory afferent did not lead to degeneration in experimental animals, if an underlying instability did not coexist 60.

The role of sympathetic postganglionic fibers in joint circulation is dual: precapillary fibers, in direct contact with blood vessels, have a vasoconstrictive function; postcapillary fibers located in proximity but not in contact with venules indirectly increase plasma extravasation by activation of the cyclooxygenase pathway and synthesis of prostaglandin, particularly

PGE2. Activation of the peripheral nervous system can initiate the major features of acute inflammation, which include vasodilation (calor and rubor), plasma extravasation (tumor), and pain sensation and a lower threshold for pain (dolor) 6i. Plasma extravasation and joint effusion, a cardinal feature of acute articular inflammation, results from the contribution of neurotransmitter release from primary afferent unmyelinated or C-fibers, sympathetic efferents, and mast cells. There is current evidence to show that the effect of

27 neurotransmitter release is dependent on the downstream presence of sympathetic terminals and mast ceils, to cause plasma extravasation. In particular, 6% adrenergic stimulation resulted in severe arthritis, as measured by plasma extravasation or chronic tissue injury. In a similar manner, adrenalectomy significantly reduce the severity of experimentally induced arthritis, illustrating a role of the neuroendocrine system in disease.

The pain of arthritis is relayed both by type IV/C PAN fibers and type HI/A3 fibers.

C-fibers are polymodal as they can respond to mechanical, thermal or chemical stimuli.

Type m/A9 fibers are mostly sensitive to mechanical or thermal stimuli. PAN can be directly stimulated by bradykinin, serotonin, and histamine or by potassium or protons, which can act to depolarize PAN. Substance P increases excitation caused by these compounds. Stimulation of small C-fibers causes release of the co-localized neuropeptides

CGRP and substance P which act synergistically to exert pro-inflammatory paracrine effects on synovium. In contrast to hormones, which are released in low concentration in circulating blood and depend on specific high affinity receptors for site-specificity, neuropeptides are delivered to the site of action by axonal transport where release is mediated by action-potential induced depolarization and calcium dependent exocytosis of vesicles. Because of these nerve ending release, the presence and density of nerve fibers is directly related to neuropeptide at that site. Furthermore, the activity of neuropeptides in a particular site is limited by the presence of peptidases, the presence of which has been documented in several articular tissues.

Of the many putative neurotransmitters released by unmyelinated primary afierents, substance P has received the most attention. Substance P is an undecapeptide that is synthesized by neurons located in the dorsal root ganglia and stored in small diameter

C-fibers nerve endings until release. These C-fibers are localized in the intima and subintima of joints, and in subchondral bone. Small C-fibers containing neuropeptides including calcitonin gene related peptide and substance P have been identified in the

28 synovium of rat, horses, rabbits. Neurokinin A and calcitonin-gene related peptides are colocalized with substance P in these nerve endings, and also have potent vasodilatory and permeability effects. Upon stimulation, release of substance P from nerve endings serve to transmit pain sensation through spinal pathways. Local antidromic release into the

interstitium and joint cavity also serves to contribute to the inflammatory reaction.

Substance P has been termed a pro-inflammatory peptide, and it increases vascular permeability, causes vasodilation (through release of nitric oxide), increases mast cells release of inflammatory mediators such as serotonin and histamine, increases leucocyte adherence, and activates lymphocytes. Substance P has a local action and its role is limited by the action of a peptidase, which is localized in the vicinity of its release. The NKl class of neuropeptide receptor has the greatest affinity for substance P. Variations in receptor density and affinity helps to explain the variable response to inflammation. Specific NKl receptor antagonists have helped our understanding of the role of substance P and may have a role in controlling the inflammatory response. Although acute inflammation is a

necessary and appropriate response to initiate repair following tissue injury, inadequate regulation of this response may lead to excessive tissue damage or chronic inflammation.

Physiological release of substance P may be protective in tissue injury to increase blood flow, synovium perfusion, and promote tissue repair. The observation of increased cellular

infiltrates in arthritic joints in areas of depleted substance P-immunoreactive fibers and

reduction in endothelial NKl receptor supports a role of neurovascular regulation to resolve chronic arthritis.62 Substance P does not appear to have a direct role in articular

permeability or blood flow.63 Increased vascular permeability, measured by extravasation

of plasma protein, was induced by intraarticular injection of bradykinin, serotonin and

histamine; CGRP potentiated the effect of these vasoactive substances to increase vascular

permeability. CGRP infusion also significantly increased blood flow to the joint.63

29 A definite role of substance P in joint pain has been demonstrated and recent treatments by the substance P depleting substance capsaicin have become available.

Capsaicin (trans-8-methyl-N-vanillyl-6-nonenamide) is the pungent ingredient in hot paprika or chili peppers. Capsaicin initially activates PAN fibers, resulting in substance P release and pain, but subsequently desensitizes or degenerates PAN fibers, suggesting a mechanisms for pain alleviation with chronic use in articular inflammation 64, Gold sodium thiomalate also causes a selective decrease in unmyelinated axons, resulting in an increased nociceptive threshold.6i A more specific role of substance P to alleviate the symptoms of arthritis has been demonstrated through immunoneutralization studies.65

NS AID are commonly used drugs in the treatment of articular inflammation. The role of prostaglandins in pain is an indirect role, as they act to sensitize PAN fibers to subsequent stimulation. There is some evidence that PGE2 and PGI can also act directly on nociceptors, but this effect is less potent than their sensitizing effect. In order to understand the effect of an inflammatory mediator or of a neuropeptide however, it is essential to remember that the result will be dependent of the density of receptors to the substance. For example bradykinin 2 receptors acutely stimulate pain in inflammation, but chronic inflammation upregulates B 1 receptors, thus reversing the hyperalgesia.66 NSAED inhibit

PGH2 synthase enzymes (also known as cyclooxgenase) and diminish the formation of

PGEl, PGE2, PGP and PGI from arachidonic acid. Corticosteroids inhibit phospholipase

A2, thus preventing the formation of arachidonate, a substrate for the cyclooxygenase and lipoxygenase pathways. At high doses, corticosteroids also inhibit D-l and TNF, which can also sensitize pain nociceptors. These mechanisms may explain the analgesic effects of

NS AID and steroids. In addition to neuropeptide release and receptor activation, degradation must be considered in our understanding of the roles of neuropeptides in pain.

Neuropeptide degradation is mediated by several amino and endo-peptidases but can also

30 be degraded by various reactive oxygen species. Several peptidases have been identified in close proximity to neuropeptide release in the joint, and these enzymes are shown to be increased in synovial tissue and fiuid of rheumatoid patients. However, the distance traveled to target cells is much greater than to post-synaptic junction complicating the net effect of neuropeptides in arthritis.67.68 Although neuropeptide Y fibers have been identified in synovial tissues including equine tissue, the role of NPY is less defined, but may be related to diminution of reflex sympathetic discharge. Finally the stress responses and hypothalamo-pituitary-adrenal axis have also implicated in inflammatory arthritis.66

The role of neuropeptides in joint disease is currently under investigation, and there appears to be differences in the contribution of neuropeptides to disease process in acute vs chronic inflammatory arthritis. In acute arthritis, loss of sensory nerves may contribute to inflanunation, as demonstrated by increased edema formation in denervated lim b s.63

Similar results have been reported in an 11-1 induced model of acute inflammation in the horse, where increased edema and decreased permeability to macromolecule were observed in denervated lim b s.7 The role of innervation in chronic arthritis is complex. Staining for

CGRP and substance P was increased in the sciatic nerve, dorsal root ganglia and periarticular tissues, but synovium staining was decreased. It appears that the role of neuropeptides in acute or chronic inflammation may vary as the distribution of sensory nerves is altered with the inflammatory response.

The therapeutic implications of the participation of neuroendocrine mechanisms in arthritis are many. Intramuscular gold or topically applied capsaicin are agents that selectively destroy type IV/C fibers, thus lowering SP levels, and have been found clinically useful. Capsaicin initially causes release of substance P from nerve ending, explaining the burning sensation felt upon initial application. Both type m/A and type IV/C fibers are stimulated by bradykinin, serotonin, histamine and prostaglandin. NS AID

31 (PGH2 or cyclooxygenase inhibitors) decrease prostanoid production, and intraarticular corticosteroids which inhibit the arachidonic acid cascade are ineffective in the treatment of inflammation and pain in arthritis.65 In addition, stimulation of primary afferent nociceptive fibers causes release of glutamate and substance P from central spinal pathways. This nociceptive input can be inhibited by stimulation of proprioceptive and tactile type I and H fibers. Stimulation of these fibers can be accomplished by high frequency low intensity transcutaneous neural stimulation, frequently used in physiotherapy.

32 Mechanisms of joint injury Cytokines Synoviocytes (D-l,n-6,TNF) Chondrocyte^ Cartilage Inflammatory degradation nriAHiaf-nrc

Hyaluronan degradation Acute joint Generation of free mjury radicals

Increased Decreased blood Neuropeptides capillary \ (CGRP, substance P) permeability

Increased Pain ____ ^ Increased intraarticular T intraarticular pressure Inunobilization volume

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54. Myers, S. L., K. D. Brandt, O. Eilam. Even low-grade synovitis significantly accelerates the clearance of protein form the canine knee. Arthritis Rheum 38:1085-1091, 1995.

55. Wallis, W., P. Simkin, W. Nelp. Protein ttraffic in human synovial effusions. Arthritis Rheum 30:57-63, 1987.

56. De Ceulaer, K., G. Balint, A. El-Ghobarey, et al. Effects of corticosteroids and local anaesthetic applied directly to the synovial vascular bed. Ann Rheum Dis 38:440-442, 1979.

57. Laurent, T. C., U. B. G. Laurent, J. R. E. Fraser. Functions of hyaluronan. Ann Rheum Dis 54:429-432, 1995.

58. Gotoh, S., J.-I. Onaya, M. Abe, et al. Effects of the molecular weight of hyaluronic acid and its action mechanisms on experimental joint pain in rats. Ann Rheum Dis 52:817- 822, 1993.

59. Vilensky, J. A., B. L. O'Connor, K. D. Brandt, et al. Serial kinematic analysis of the canine knee after L4-S1 dorsal root ganglionectomy: Implications for the cruciate deficiency model of osteoarthritis. J Rheumatol 21:2113-2117,1994.

60. O'Connor, B. L., M. J. Palmoski, K. D. Brandt. Neurogenic acceleration of degenerative joint lesions. J Bone Joint Surg 67-A:562-569, 1985.

61. Basbaum, A. L, J. D. Levine. The contribution of the nervous system to inflammation and inflammatory disease. Can J Physiol Pharmacol 69:647-651, 1991.

37 62. Walsh, D. A., M. Salmon, P. I. Mapp, et al. Microvascular substance P binding to normal and inflamed rat and human synovium. J Pharm ExpTher 267:1993.

63. Cambridge, H., S. D. Brain. Calcitonin gene-related peptide increases blood flow and potentiates plasma protein extravasation in the rat knee. Brit J Pharmac 106:746-750, 1992.

64. Colpaert, F. C., J. Donnerer, F. Lembeck. Effects of capsaicin on inflammaiton and on the substance P content of nervous tissues in rats with adjuvant arthritis. L^e Sci 32:1827-1834, 1983.

65. Konttinen, Y. T., P. Kemppinen, M. Segerberg, et al. Peripheral and spinal neural mechanisms in arthritis, with particular reference to treatment of inflammation and pain. Arthritis Rheum 37:965-982, 1994.

66. Stemberg, E. M., G. P. Chrousos, R. L. Wilder, et al. The stress response and the regulation of inflammatory disease. Ann Intern Med 117:854-866, 1992.

67. Lowe, J. R., J. S. Dixon, J. A. Guthrie, et al. Seram and synovial fluid levels of angiotensin converting enzyme in polyarthritis. Ann Rheum Dis 45:921-924, 1986.

68. Sreedharan, S. P., E. J. Goetzl, B. Malffoy. Elevated synovial tissue concentration of the common acute lymphoblastic leukaemia antigen (CALLA)-associated neutral endopeptidase in human chronic arthritis. Immunology 71:142-144,1990.

38 CHAPTER 3

PRESSURE-VOLUME RELATIONSHIPS IN EQUINE MIDCARPAL

JOINTS Introduction

Normal intra-articular pressure (TAP) is subatmospheric in most joints placed at an angle of ease.i Maintenance of normal lAP is important for optimal synovial blood flow and transsynovial fluid exchanges. Joint effusion, a common clinical finding in joint disease, can lead to increased lAP and decreased synovial membrane blood flow.zj

Decreased synovial membrane blood flow may result in altered synovial fluid filtration and composition, articular metabolite accumulation, hypoxemia, and acidosis, all of which can contribute to altered chondrocyte metabolism and promote further articular injury. 2-6

Parameters that have been used to describe the pressure volume (PV) relationship of joints include elastance [change in pressure per unit change in volume (AP/AV)] and its reciprocal, compliance [change in volume per unit change in pressure (AP/AV)] .7.8

Elastance is a measure of resistance to deformation, and compliance reflects the ability of a structure to deform. Other useful parameters are hysteresis is a measure of the difference in behavior of elastic tissues during progressive increased or decreased stress, caused by a greater rate of disengagement of wall elastic elements during unloading than the rate of recruitment during loading.^ Stress-relaxation is a measure of the decay of pressure over time, under a constant volume or strain, caused by relaxation of the joint capsule.

However, because fluid absorption from the joint may contribute to the observed decay in

39 pressure, the expression "relaxation of lAP" will be used in the present study, since fluidabsorption was not measured.

Numerous studies have reported on the PV relationships of joints.i-2.7-i3 These studies have concluded that intra-articular volume, joint capsule compliance, joint angle, joint measurement, site measurement, history of prior distention, and joint activity are important determinants of IAP.8 More specifically, increased intra-articular volume, decreased capsular compliance caused by thickening and fibrosis, and decreased joint angle

(i.e., flexion) contribute to increased lAP. Different joints have different baseline pressures. 14 Compartmentation within a joint has been demonstrated in the rabbit knee and is known to alter the pressure response to fluid infusion through uneven distribution of the infusate.i 1 There is net transsynovial filtration during prolonged immobilization that results in a mild increase in intra-articular volume and pressure. 15 Periarticular muscle contraction during locomotor activity increases lAP particularly in the presence of joint effusion. All of these studies have substimted polyionic isotonic solutions or low-viscosity nonabsorbable oil for synovial fluid to describe these phenomena. Furthermore, in some studies, the skin that covers the joint, a major supporting structure, was removed, which may alter the compliance of the joint.2.9

The characteristics of the PV relationships of an organ can be divided into two separate parts^: 1) the elastic forces of the tissuei^ and 2) the elastic force caused by surface tension of the fluid that lines the inside of the organ, t? Synovial fluid is an ultrafiltrate of plasma to which hyaluronate has been added. Hyaluronate is a high-molecular-weight proteoglycan that imparts a high viscosity to synovial fluid.

Hyaluronate is also the principal glycosaminoglycan of the interstitium. The viscoelastic non-Newtonian behavior of synovial fluid may yield different PV relationships than that obtained with saline or other fluids, In addition, the use of synovial fluid abolishes the

40 effect of surface tension that would be present at the interface of two different fluids, as when saline or oil are used, and allows for more accurate determination of the elastic properties of the joint capsule. The boundary lubricant properties of the hyaluronan molecule, which forms an intimate bond with synoviocytes, may also alter the mechanical properties of the joint walls.

The equine midcarpal joint is the most common joint affected by traumatic injury in performance animals. Traumatic injury to the midcarpal; joint is accompanied by effusion and lameness. Clinically evident effusion probably results in increased lAP, which may in effect reduce synovial membrane blood flow. The equine midcarpal joint has been used in many models of articular disease for mechanistic or therapeutic investigations. 19 To improve our understanding of the pathogenesis of acute or chronic joint injury and the effect of joint effusion on blood flow, it is important to investigate the normal relationship between synovial fluid volume and LAP within an intact joint.

The purpose of the present study was to describe the PV relationships of the equine midcarpal joint and to determine the effect of joint angle, nature of infiisate, and prior distention on these relationships.

Materials and methods

j-Twenty-four healthy adult horses (350-500 kg) and 48 midcarpal joints were used for this study. Criteria for inclusion of horses in the study included carpal joints free of disease on the basis of physical examination and gross examination of the articular cartilage and synovium at postmortem. An approved Institutional Laboratory Animal Care and Use protocol was obtained before this study was begun.

Anesthesia-HoTses were sedated with xylazine (0.5 mg/kg iv), anesthetized with 5% guaifenesin (intravenously to effect) and thiamylal (4 mg/kg iv), and maintained with

41 pentobarbital sodium (5-15 mg • kg-i • h-i iv). All horses were intubated with a cuffed endotracheal tube, positioned in dorsal recumbency, and mechanically ventilated with

100% oxygen. A 20-gauge catheter was inserted into a facial artery for direct measurement of arterial blood pressure and collection of samples for determination of arterial pH and blood gasesa. Systolic, mean, and diastolic arterial blood pressures were continuously displayed, and arterial pH and blood gases were determined hourly. Drug administration and mechanical ventilation were adjusted to maintain the mean arterial pressure at > 70 mmHg, arterial PO 2 at > 150 Torr, and arterial PCO 2 at < 60 Torr.

Instrumentation-Thc horses were positioned in dorsal recumbency, and both forelimbs were suspended from a rack by a rope tied around the pastern. Tension was not exerted on the limb. The midcarpal joints were set at the predetermined angle (90° or 135°) with a goniometer (Fig. 3.1). A 20-gauge needle connected to a stiff-walled polyethylene extension tubing filled with heparinized saline and closed by a three-way connector was inserted into each of the medial and lateral dorsal midcarpal joint pouches and the medial and lateral palmar midcarpal joint pouches. Each connector was attached to a horizontally positioned pressure transducer^ placed at the same height as its corresponding needle, and each pressure transducer was then calibrated with a mercury manometer before measurement of LAP. Total compliance of the system was 0.45 |il/mmHg between 0 and

100 mmHg. Transducer drift was 0.008 mmHg/minc. The dorsal midcarpal joint was also cannulated with a 20-gauge needle placed through the extensor carpi radialis tendon for infusion and withdrawal of fluid. A calibrated pumpd adjusted to deliver a rate of 2.47 ml/min was used for infusion and withdrawal of fluids. This rate of fluid transfer was chosen to minimize artifact caused by relaxation of lAP of the joint capsule during infusion while allowing enough time to measure small pressure changes. All pressure tracings were

42 continuously displayed and recorded on a physiography. Synovial fluid was obtained from the tarsocrural joints of the same horse and used on the same day. Synovial fluid was considered to be normal based on color, clarity, and viscosity. The viscosity of synovial fluid obtained from normal equine tarsocrural joints has been reported. 18

Protocol

Eight experiments were conducted (n = 6 midcarpal joints per experiment). Only one experiment per joint was performed.

Experiments 1-4: effect o f joint angle and infusate. The PV relationships in the midcarpal joint were determined at a joint angle of 90° by infusion of synovial fluid {Expt I) or isotonic saline (0.9% NaCl) (expt 2), or at a joint angle of 135° by infusion of synovial fluid (expt 3) or isotonic saline (expt 4).

Experiments 5 and 6: history dependence ofPV curves. The effect of sequential distention on articular compliance at lAPs in the low (< 30 mmHg) or high (30-80 mmHg) range was determined by continuous infusion at the usual rate of synovial fluid to an lAP of 30 mmHg followed by withdrawal at the same rate. This cycle was repeated twice more, then lAP was increased from 30 to 80 mmHg followed by withdrawal to return to 30 mmHg; similarly, this cycle was repeated twice more (expt 5). In six additional joints {expt 6), lAP was first increased from 30 to 80 mmHg for three cycles, followed by three cycles at low range (up to 30 mmHg).

Experiments 7 and 8: relaxation oflAP over time. Relaxation of lAP was measured at angles of 90° {expt 7) and 135° {expt 8) by continuous infusion of synovial fluid in increments of 20 mmHg, up to 80 mmHg. After each 20-mmHg increase in LAP, the infusion was stopped and the pressure decay was measured for 5 min before the next increment.

43 Analyses

Ali pressure tracings were analyzed by scanning each tracing into a software program^ to obtain data points. These data points were used to generate PV curves for each subject and each experiment. These curves were analyzed with linear and nonlinear regression.

Experiments 1-4. Examination of the PV curves revealed a sigmoid relationship that was best fitted to an exponential equation (R 2 > 0.98), as previously described.? Each PV curve was fitted to the exponential equation LAP = A x e(B x volume) _ C, where B is the fractional change in pressure per unit change of volume, thus defining the curvature of the relationship, and A and C on constants. The term B was used for statistical analysis in testing for the effect of joint angle, fiuid infused, hysteresis, and compartmentation. All parameter values were obtained by an iterative algorithm by using the Levenberg-Marquardt method, with R2 > 0.98. Elastance was calculated for each increment in pressure and volume (AP/AV) and was plotted against mean lAP. This lAP/V vs. lAP relationship was linear below or above atmospheric pressure, and the slope of the linear regression equation was compared at pressure ranges below and above atmospheric pressure. Hysteresis was computed by measuring the areas under the infusion and withdrawal curves^ and calculated as percent [(area under infusion - area under withdrawal)/area under infusion].

Experiments 5 and 6. PV relationships were linear below and above 30 mmHg (R 2 >

0.99). Linear-regression equations of the form lAP = a + b x volume where a is the intercept and b is the slope AP/AV, were fitted to the low (baseline to 30 mmHg) or high

(30-80 nunHg) PV curves by using the least squares method. The slope of each curve, representing elastance, was used for statistical analysis.

44 Experiments 7 and 8. Articular relaxation at a given pressure was best described as a logarithmic decaying function of time.9 The relationship P, = ?t - s In t, where ?t is pressure at a given time t, Pj is initial pressure, and s is slope of the curve, was determined for each relaxation of the lAP curve at each 20 mmHg increment. The slopes of each stress relaxation curve were compared among incremental pressures for each joint angle.

Statistical Analysis

Student's t-test was used to compare baseline lAPs between the two joint angles. A four-way fixed-model analysis of variance was used to compare the B coefficients of the

PV curves, with joint angle, infusate, joint compartment, and infusion of withdrawal as the independent variables. Body weight of the horses was tested and found not to correlate with the data, so it was not included as a covariate. A four-way repeated-measure analysis of variance was used for comparing the linear slopes for joint angle, compartment, and low or high pressure, with cycles as the repeated measure. A two-way analysis of variance was used to compare the hysteresis ratio between infusâtes (saline or synovial fluid) and joint angle (90° or 135° angle). A three-way repeated-measure analysis of variance was used to compare the relaxation of lAP function between joint angles and among pressure increments, with slope as the repeated measure. For all analyses, when a main effect or an interaction was detected, a Newman-Keuls post hoc comparison test was used to identify the different means. Significance was set at P < 0.05, and data are expressed as means ±

SE.

45 Results

Baseline lAPs measured -2.7 ±0.1 mmHg [95% confidence interval (Cl) = -4.84, -0.5] at

135° and 0.3 ± 0.1 mmHg (95% Cl = -2.18,2.44) at 90° midcarpal joint angle. lAP was significantly higher (P = 0.0062) at 90° than at 135° angle.

Description ofPV Relationship in Midcarpal Joints

The PV relationship in equine midcarpal joints was sigmoid and was best defined with two exponential equations (Fig 3.2 and 3.3): for subatmospheric pressures: lAP =-7.20 Xe(-3.46x volume)-3.61 (1) and for supra-atmospheric pressures lAP = 40.74 X g (0 . 11 X volume) -35.46 (2)

Elastance (E) was linearly related to lAP below or above atmospheric pressure (Fig. 3.4).

The regression equation below atmospheric pressure was

E = 19.13 - 3.276 X lAP (/?2= 0.90, P = 0.0008), (3) and that above atmospheric pressure was

E = 4.38 + 0.108 X lAP (/?2 = 0.99, P < 0.0001). (4)

Examination of the slopes of the regression equations (3.27 vs. 0.0108) demonstrates the much greater resistance to distention of the joint (greater elastance, lesser compliance) at subatmospheric than supra-atmospheric pressures.

In the present study, we were unable to demonstrate significant compartmentation of fluid during infusion or withdrawal, indicating that in dorsal recumbency for joint angles of 90° or 135° angle, fluid was distributed evenly between all four joint pouches.

46 Hysteresis was detected for all joint angles for synovial fluid or saline infusion

(Fig. 3.5, A and B). Hysteresis at 135° angle was 38 ± 2 and 22 ± 2% for synovial fluid and saline infusion, respectively. Hysteresis at 90° was 57 ± 3 and 30 ± 2% for synovial fluid and saline infusion, respectively. There was significantly greater hysteresis when synovial fluid was used as the infusate with a joint angle of 90° (P < 0.005).

In two horses, rupture of the joint capsule with subcutaneous leakage of synovial fluid was observed at a joint angle of 135° and lAP of 80 mmHg. The rupture occurred in the lateral palmar articular joint pouch.

Effect of Joint Angle

Decreased joint angle from 135 to 90° significantly increased subatmospheric (P =

0.028) and supra-atmospheric (P = 0.0016) articular elastance (Figs. 3.2 and3.3).

Effect o f Type o f Infusate

Infusion of saline resulted in a significantly higher articular elastance than that measured with infusion of synovial fluid (Fig.3. 6) at 135° (P = 0.045) and a trend toward higher elastance at 90° (P = 0.063).

Effect o f Prior Distention on PV Relationships

Sequential distention cycles (history dependence) did not significantly affect PV relationships at lAPs < 30 mmHg. At high lAPs (30-80 mmHg) a significant increase in elastance was observed at the second and third distention cycles, when the high-pressure cycles followed low-pressure cycles (P < 0.(X)01) (Fig.3.7). Elastance at lAPs < 30 mmHg was 8.00 ± 0.78 mmHg/ml and was not different between cycles. Elastance at the first high-pressure cycle after three low-pressure cycles was 14.87 ± 1.00 mmHg/ml, and values for subsequent high-pressure cycles were 17.09 ± 1.25 and 17.27 ± 1.24

47 mmHg/ml. There was a trend toward an effect of joint angle on elastance for low or high lAP (P = 0.087).

Relaxation oflAP

Sequential increases in lAP by 20-mmHg increments allowed to better define elastance at supra-atmospheric pressures ranging from 0 to 20,20 to 40,40 to 60, and 60 to 80 mmHg. There was a significant difference (P < 0.005) between all incremental pressures, and the lowest pressure range was associated with the lowest elastance (greater compliance) (Fig. 3.8). A significant relaxation of lAP was observed when pressures were increased to 40,60, or 80 mmHg. At 20 mmHg, relaxation of lAP was minimal for that measurement period. The slope of the stress relaxation curve of the pressure-time relationship was a positive logarithmic function of lAP (Fig. 3.9).

Discussion

The PV relationship of the equine midcarpal joint is a characteristic sigmoid curve that is best described by two exponential equations, one for subatmospheric pressures and one for supra-atmospheric pressures. This dual relationship reflects the difference in compliance observed at sub- or supra-atmospheric pressures, indicating that the joint capsule resists distention at normal (subatmospheric) pressures; however, after increases in intra-articular volume, as occurs with effusion, there is relaxation of the joint capsule to accommodate the increased volume. Examination of the relationship between elastance and

LAP better illustrates this relationship, as the regression coefficient is much greater at subatmospheric than supra-atmospheric than supra-atmospheric pressures (3.27 vs.

0.108/ml). This sigmoid PV relationship has been described in other species^ and is comparable to that observed in the interstitial space, supporting the statement that the joint cavity, because of its lack of a basement membrane, is an expansion of the interstitial

48 space.20.2i The low compliance observed at subatmospheric pressure may help prevent edema formation and help maintain a normal negative lAP, which favors fluid exchanges.

The more compliant joint observed at supra-atmospheric pressures up to 30 mmHg may help prevent impaired blood flow and preserve synovial membrane metabolism as well as decrease joint pain. The decreased compliance at higher pressure may impede net filtration of fluid and prevent further effusion.

In our study, compartmentation of fluid within the different midcarpal joint pouches was not observed. This finding contrasts with that observed in the rabbit stifle (knee) jointii and may reflect the non-weight-bearing position of the leg, the larger joint compartments of the horse, or the difference in the anatomy of the stifle joint vs. the midcarpal joint. This finding validates the future use of the dorsal joint compartments in studies that investigate lAP of the equine midcarpal joint in the non-weight-bearing position. However, we found that the palmar joint pouches were more difficult to cannulate because of their smaller size, particularly in smaller subjects.

We observed that the response to volume infusion was different when saline was used instead of synovial fluid, with saline infusion resulting in a lower joint compliance than when synovial fluid was used. Another study reported increased compliance with a polyionic solution compared with a nonabsorbable oil.? The viscosity of synovial fluid is given by its high hyaluronate content. This high-molecular-weight molecule has properties that are shear-rate dependent, i.e., it is more viscous at low shear rates. In addition, synovial lining cells have receptors for hyaluronate, and the cell-hyaluronate unit may yield different properties to the joint capsule than each component taken separately .2 2

In the present study, we measured significant hysteresis, reflecting a difference in the PV relationship between volume infusion or withdrawal. Compared with saline, hysteresis was significantly greater when synovial fluid was used as the infusate. This

49 observation is similar to that observed in the lung when saline is used instead of surfactant.

In the lung, saline inflation markedly decreases hysteresis in the absence of surfactant; with surfactant, normal hysteresis is restored.23 The high-molecular-weight hyaluronate molecule may play a similar role in the joint, where, by preventing joint collapse, it helps maintain subatmospheric pressure, which favors fluid exchanges. Furthermore, it is doubtful that all hyaluronan molecules are initially removed when saline or oü are used as infusate in PV studies. It is more likely that they are progressively washed out of the joint with each PV loop. Therefore, when fluids other than synovial fluid are used there is a fluid-synovial fluid interface, the composition of which changes with each cycle of infusion or withdrawal. Our findings suggest that when fluids other than synovial fluid are used, the mechanical properties of the hyaluronan-synoviocyte unit are altered; perhaps this illustrates the important boundary properties of hyaluronan.

As previously described,? joint flexion (135 to 90° angle) significantly decreased articular compliance, probably by increasing tension on the joint capsule. This finding may explain the clinical observation that with severe effusion most horses will resent joint flexion and prefer to hold their joint at ~ 135°.

In the present study, increased elastance or decreased compliance was measured after the first infusion cycle at high lAP. The total compliance of the joint is the sum of the compliance of each part. The decreased compliance observed with subsequent cycles at high pressure may reflect distribution of fluid within the joint during the first cycle or may be related to the relatively rapid rate of infusion. Normal human and dog lungs undergo a decrease in compliance over time when ventilated spontaneously or mechanically.24 In a previous study by Knight and Levick,? compliance increased with subsequent infusions, in contrast to our findings. Knight and Levick attributed this increased compliance to plastic deformation of the joint capsule. In their study, infusion rate was slower than our rates. In

50 addition, the skin overlying the joint had been removed, which may have changed the mechanical properties of the periarticular tissues.

Relaxation of lAP was significant, particularly at higher lAPs. In our study, stress- relaxation was a better function of the natural log of time than a linear function, which suggests that with prolonged observation a non-zero asymptote was being approached.9

Relaxation of lAP is a measure of pressure decay under constant volume or strain.

Alternative explanations for the decay in lAP include fluid absorption from the joint space, slow change in joint angle, and slow fluid distribution within the joint space.? Calculation of relaxation of lAP at 80 nunHg gives a value of 7.56 mmHg/5 min (see equation in Fig.

9 legend). Compliance (I/elastance) at 80 mmHg was 0.045 ml/mmHg (see Fig. 5), such that absorption of 0.225 ml (225 fil) of synovial fluid in 5 min would have been necessary to decrease lAP by 7.56 mmHg. Hydraulic conductance of synovial fluid in the equine midcarpal joint at normal or increased lAP is unknown. In other species, the rate of turnover of synovial fluid estimated by three different methods (rate of change in pressure, protein influx, and protein efflux) yielded values ranging from 1.3 to 6.7 p.l/cm2 of synovium per hour for normal joints; however, these values probably are different with increased lAP, considering that hydraulic conductance for Ringer solution was increased almost sixfold with lAP > 9 and < 25 cmHzO. The effect of increased lAP on synovial membrane blood flow and clearance also needs to be considered. We have shown that blood flow in synovial membrane of equine midcarpal joints (joint angle of 135°) was decreased from 108 ± 36 to 12± 7 and 11 ±3 p.lmin-i-g-i atlAPs of 30 and60 nunHg, respectively. Such low blood flow could alter fluid clearance from the joint cavity such that extrapolation of hydraulic conductance values obtained at 25 cmHzO may not be valid.

Therefore, we cannot conclude on the contribution of fluid absorption from the joint to the decrease in lAP over time. In our study, horse limbs were secured and fixed in position

51 for a minimum of 1 h before the start of the study. Our study did not demonstrate compartmentation in the midcarpal joint Infusion of fluid was associated with an immediate increase in lAP in all four compartments. Therefore, it is unlikely that fluid distribution would be responsible for the decay in pressure over time. In our study, we used a short measurement period (5 min) to measure the rate of pressiu'e decay. This short measurement period allowed us to use a simple logarithmic transformation to linearize the pressure-time relationship. The characteristics of biomechanical relaxation of tissue is a function of the length of the experiments, and long-duration experiments are necessary to accurately describe this phenomenon.^ These experiments are mathematically more complex, and therefore were not performed. Our goal was to describe the short-term effect of time on lAP, and we demonstrated that in future studies, particularly at high lAPs, measurements should be made immediately after pressure increases to minimize the effect of relaxation of lAP.

In conclusion, our study described the PV relationships of the equine midcarpal joint and highlighted the importance of joint angle, nature of infusate, and time on these relationships. Our findings will serve as a useful basis for future studies that investigate articular PV relationships.

Footnotes a ABL 500, Radiometer, Copenhagen, Denmark b P23 XL Statham transducer, Gould, Cleveland, OH c Personal communication from Omega Corp., Cleveland, OH d Model 940 infusion-withdrawal pump; Harvard Apparatus, Dover, MA e Model 7D Polygraph, Grass Instruments, Quincy, MA f Flexitrace, Tree Star, Campbell, CA

52 Figure 3.1. Illustration of forelimb of horse, demonstrating position of needles for measurement of intra-articular pressure in midcarpal joints.

53 Joint Angle 90° or 135

To physiograph To physiograph

54 Figure 3.2. Pressure-volume relationships in equine midcarpal joints demonstrating sigmoid shape of curve and difference between sub- and supra-atmospheric pressures.

Elastance is significantly lower (greater compliance) at 135° than 90° angle (P=0.0016).

55 Intraarticular pressure (mm Hg) -*i\)w*»(no)N00(O ooooooooooo

o> -

09 “ % Angle=90

Average Figure 3.3. Pressure-volume relationships in equine midcarpal joints at subatmospheric

(normal ) pressures. Elastance is significantly lower (greater compliance) at 135° than 90° angle (P=0.028).

57 Intraarticular pressure (mm Hg)

o <30

% CO

59 45 40 |>3S ^ 3 0 I 25 8 20 # 1 5 w 10 5 0 -10 0 10 20 30 40 50 60 70 80 Intraarticular pressure (mm Hg)

60 Figure 3.5. PV relationships in equine midcarpal joint during infusion or withdrawal of saline or synovial fluid at joint angles of 90° (A) or 135° (B). Hysteresis calculated as (area under infusion curve-area under withdrawal curve)/area under infusion (in %), is significantly greater with synovial fluid than saline (?<0.005) and at 90° than at 135° angle

(P<0.005).

61 Intraarticular preaaura (mm Hg) CO Intraarticular pressure (mm Hg) 8 o S o

2 c o> t

o o

fO to I Figure 3.6. PV relationships in equine midcarpal joints at 135° infused with isotonic saline or synovial fluid. Elastance was significantly greater with saline than with synovial fluid (P=0.045).

63 Intraarticular pressure (mm Hg) 0 3 en 00 o o o

ro-

0 0 -

O ■ to < ' 5' c a Figure 3.7. A: PV relationships at 3 low-pressure cycles (<30 mmHg) followed by 3

high-pressure cycles (30-80 mmHg) and at 3 high-pressure cycles followed by 3 low-

pressure cycles. B: inset of A; shows that elastance significantly increased at high-pressure

cycles after prior distention at low pressures.

65 99

Intraarticular pressura (mm Hg) ^ o>Ai>oUi>kUio)'^oB«e o o o o o o o o o

ro Ir - î » j

o> -

Intraarticular pressure (mm Hg) Ê S S S S § m

o*_i Figure 3.8 Relaxation of lAP as a function of natural log of time for 20-mmHg increments of lAPS (joint angle 135”). Slope of lAP-relaxation (lAP-R) curve is significantly increased with increasing lAP (P< 0.005).

67 80- 8=4.83

? 7 0 . E Initial lAP = 80 mm Hg £ 6 0 - 8=3.96

3 50- Initiai lAP = 60 mm Hg 8=2.56 &| 40- Initial lAP = 40 mm Hg I 30- u 8=0.69 r 20- Initial lAP s 20 mm Hg 1 , 0.

-2.5 -2 -1.5 -1 -0.5 0 0.5 Ln time (min)

68 5 -,

3U54 « 3 I 254 I #154 S Q. — 0 .5-

2.8 3 3.2 3.4 3.6 3.8 4 4.2 4.4 Ln lAP (mm Hg)

Figure 3.9 lAP-R as a function of lAP. There is a significant increase in slope of lAP-R curve with increased initial lAP. LAP = -8.37 + 3.00 x In (lAP) (R2 = 0.99, P < 0.0001).

69 References

1. Levick, J. R. An investigation into the validity of subatmospheric pressure recordings from synovial fluid and their dependence on joint angle. J Physiol (ùnd) 289:55-67, 1979.

2. Geborek, P., K. Forslind, F. A. Wollheim. Direct assessment of synovial blood flow and its relation to induced hydrostatic pressure changes. Ann Rheum Dis 48:281-286, 1989.

3. Hardy, J., A. L. Bertone, W. W. Muir. Determination of synovial membrane blood flow in equine joints using colored microspheres. Abstract, Vet Surg 22:383, 1993.

4. Clark, C. C., B. S. Tolin, C. T. Brighton. The effect of oxygen tension on proteoglycan synthesis and aggregation in mammalian growth plate chondrocytes. J Orthop Res 9:477-484, 1991.

5. Lane, J. M., C. T. Brighton, B. J. Menkowitz. Anaerobic and aerobic metabolism in articular cartilage. J Rheumatol 4:334-342, 1977.

6. Levick, J. R. Hypoxia and acidosis in chronic inflammatory arthritis; relation to vascular supply and dynamic effusion pressure. J Rheumatol 17:579-582, 1990.

7. Knight, A. D., J. R. Levick. Pressure-volume relationships above and below atmospheric pressure in the synovial cavity of the rabbit knee. J Physiol (Lond) 328:403- 20, 1982.

8. Levick, J. R. Joint pressure-volume studies: their importance, design and interpretation. J Rheumatol 10:353-7, 1983.

9. Knight, A. D., J. R. Levick. Time-dependence of the pressure-volume relationship in the synovial cavity of the rabbit knee. J Physiol (Lond) 335:139-52,1983.

10. James, M. J., L. G. Cleland, A. M. Rofe, et al. Intraarticular pressure and the relationship between synovial perfusion and metabolic demand. J Rheumatol 17:521-527, 1990.

11. Knight, A. D., J. R. Levick. Physiological compartmentation of fluid within the synovial cavity of the rabbit knee. J Physiol (Lond) 331:1-15, 1982.

12. Levick, J. R. The influence of hydrostatic pressure on trans-synovial fluid movement and on capsular expansion in the rabbit knee. J Physiol (Lond) 289:69-82, 1979.

13. Levick, J. R. Flow through interstitium and other fibrous matrices. Q J Exp Physiol 72:409-37, 1987.

14. Simkin, P. A., R. S. Benedict. Hydrostatic and oncotic determinants of microvascular fluid balance in normal canine joints. Arthritis Rheum 33:80-86, 1990.

70 15. Levick, J. R. Synovial fluid and trans-synovial flow in stationary and moving normal joints. In: Helminen HJ, Kiviranta I, Tammi M, Saamanen A-M, Paukkonene K, Jurvelin J, ed. Joint loading. Bristol; Wright; 1987; 148-186.

16. Desjardins, M. R. Cartilage healing: a review with emphasis on the equine model. Can VetJ 31:565-572, 1990.

17. Guyton, A. C. Pulmonary ventilation. In: Guyton AC, ed. Textbook of medical physiology. Philadelphia; W B Saunders Co; 1991;.

18. Korenek, N. L., F. M. Andrews, J. M. Maddux, et al. Determination of total protein concentration and viscosity of synovial fluid from die tibiotarsal joints of horses. A m J Vet Res 53:781-784, 1992.

19. Jones, D. L., S. M. Barber, C. E. Doige. Synovial fluid and clinical changes after arthroscopic partial synovectomy of the equine middle carpal joint. Vet Surg 22:524-530, 1993.

20. Guyton, A. C. The body compartments: extracellular fluids; intracellular fluids; interstitial fluid and edema. In: Textbook of medical physiology. Philadelphia; W B Saunders Co; 1991;.

21. Hadler, N. A. The biology of the extracellular space. Clin Rheum Dis 7:71-97, 1981.

22. Johnson, B. A., G. K. Haines, L. A. Harlow, et al. Adhesion molecule expression in human synovial tissue. Arthritis Rheum 36:137-146, 1993.

23. Guyton, A. C. Pulmonary ventilation. In: Textbook of medical physiology. Philadelphia; W B Saunders Co; 1991; 402-413.

24. Ferris, B. G., D. S. Pollard. Effect of deep or quiet breathing on pulmonary compliance in man. J Clin Invest 39:143-147, 1960.

71 CHAPTER 4

DETERjVHNATION OF SYNOVIAL MEMBRANE BLOOD FLOW IN EQUINE JOINTS BY COLORED MICROSPHERES AND EVALUATION OF THE EFFECT OF INCREASED INTRAARTICULAR PRESSURE

Introduction

The synovial membrane functions to maintain articular homeostasis by providing a pathway for the exchange of nutrients and metabolic by-products between blood and synovial tissues, including articular cartilage h2. Optimal oxygen delivery to articular tissues serves to maintain normal synovial fluid composition 3,4 chondrocyte metabolism, and ensure normal matrix composition and turnover 5-7. The efficiency of exchange between the synovial membrane capillaries and joint cavity is dependent on capillary density, capillary depth and blood flow 4.

Previous investigations have used indirect methods such as clearance of small solutes or radioactive tracers to measure blood flow to articular tissues 8,9. These methods provide a relative measure of alterations in flow and do not account for the heterogeneity in blood flow distribution among different articular tissues. Direct quantitation of blood flow to the synovial membrane of normal joints may provide a better understanding of the local influence of the circulation on joint health. Radioactive microspheres were used in a study that compared blood flow in hard and soft tissues of the radiocarpal and stifle joints of normal dogs at rest and during treadmill exercise to. That study illustrated the usefulness of the microsphere technique to quantify blood flow in tissues, but had the inconvenience of

72 using radioactivity. Colored microspheres provide a quantitative, safe and relatively simple method of blood flow measurement for use in small tissue samples, and have been used to measure blood flow in several different organs, including the heart and gut

Indirect methods of blood flow measurements have helped to define the determinants of synovial membrane blood flow to be defined, which include local temperature, vasoactive substances, and intraarticular pressure. In turn, intraarticular pressure is dependent on synovial fluid volume, joint angle and joint capsule elastance. In chronic inflammatory arthritis, current evidence suggests that a state of hypoxic acidosis exists, and that the worst hypoxia and acidosis occurs in joints with advanced fibrosis, marked effusion and poor blood flow 3,4,14,15. Furthermore, flexion or usage of inflamed joints exacerbates increases in intraarticular pressure, impairs blood flow, and aggravates hypoxia 14. Intraarticular pressures of up to 170 mmHg have been measured in rheumatoid knee joints of human patients submitted to isometric quadriceps contractions In these rheumatoid patients, oxygen partial pressures fell significantly during exercise. Significant reactive hyperemia occurred after exercise and evidence of reperfusion injury was demonstrated by an increase in lipid peroxidation products in synovial fluid of exercised knees, and by an increased formation of fluorescent monomeric and polymeric IgG 16.

Significant hypoxic acidosis was measured at intraarticular pressures greater than 45 mmHg 14. Hypoxic acidosis can contribute to articular inflammation by impairment of synovial membrane and chondrocyte metabolism I7,is, lysosomal enzyme release, acceleration of inflammatory processes, and synovial membrane proliferation 9. Therefore synovial membrane blood flow plays a critical role in the pathogenesis of chronic arthritis.

Little is known of the influence of these determinants in normal joints. Determination of blood flow and blood flow distribution to normal joints would allow comparison to blood flows in acute or chronic articular disorders. Knowledge of the influence of the articular

73 environment, including joint angle, intraarticular pressure, temperature, innervation and vasoactive agents on blood flow to the joint would provide a better understanding of the dynamic events that occur during joint disease 8.

In the present study, the synovial membrane was defined as that tissue consisting of an intimai layer or synovium, 1-5 cells in depth, adjacent to the joint cavity, containing a rich capillary plexus and deeper lymph vessels 8.19, and a subintimal or subsynovial layer, principally composed of fibrous, areolar or adipose tissue depending on the function and location within the joint as originally proposed by Key 20. Morphometric studies have demonstrated that areolar synovial membrane has the greatest capillary density, and that the capillary density is greatest within 25 p.m of the synovial surface 8 .21. it is thought that this most superficial capillary plexuses of the synovial membrane contributes to the majority of solute exchanges 8.

The purpose of the present study was to quantify the normal regional distribution of synovial membrane blood flow in the horse using colored microspheres, and to determine the effect of acute increases in intraarticular pressure on synovial membrane blood flow.

Material and methods

Horses - Six healthy adult horses (350-500 kg) and 8 forelimbs were used for this study.

Criteria for inclusion of horses in the study included midcarpal and metacarpophalangeal joints free of disease based on physical examination and gross examination of the articular cartilage and synovial membrane at post-mortem. An Institutional Laboratory Animal Care and Use protocol was approved for this study.

Experimental design and protocol - Blood flow to the synovial membrane of the dorsal antebrachiocarpal; dorsal, lateral palmar and medial palmar midcarpal; and dorsal and

74 palmar metacarpophalangeal joints was determined Aom a randomly selected foreiimb in all horses (n=6), and from the contralateral limb of 2 horses. The fibrous capsule of the dorsal pouch of the midcarpal joint (n=8) was also harvested for blood flow determination and comparison with synovial membrane blood flow at the same site. One forelimb from each horse (Group 1, n=6) was used for blood flow determination at baseline (trial 1) and following increased midcarpal joint intraarticular pressures of 30 (trial 2) and 60 mmHg

(trial 3). In 2 horses, the contralateral limb (Group 2; n=2) was instrumented and blood flow was determined at corresponding times (Trials 1,2, and 3) but without increased midcarpal joint intraarticular pressure.

Horses were allowed to stabilize under general anesthesia for 1 hour before each experiment. Drug administration and mechanical ventilation were adjusted to maintain mean arterial pressure > 70 mmHg, PaOi > 150 mmHg and PaCO? < 60 mmHg. After 1 hour, a randomly selected forelimb was suspended from a rack, which placed the midcarpal and metacarpophalangeal joints at approximately 40 cm and 64 cm above the level of the right atrium, respectively. The limb was positioned to produce a 1350 angle between the radius and third metacarpus (verified with a goniometer) and instramented with catheters for microsphere determination and needles and pressure transducers for intraarticular pressure measurement and synovial fluid infusion (Rg I). Regional blood flow to the selected forelimb (Group I) were determined at baseline (trial 1) and following increased midcarpal joint pressures of 30 mmHg (trial 2) and 60 mmHg (trial 3) with the use of a different microsphere color for each trial. Intraarticular pressure was increased by intraarticular infusion of synovial fluid collected from the tarsocrural joint of the same horse. To counter for stress-relaxation 22, intraarticular pressure was maintained at the predetermined value by continuous infusion of synovial fluid for the duration of blood flow measurement (fixed pressure, variable rate of infusion). Intraarticular pressure was allowed to return to baseline

75 for 10 minutes before each increase in intraarticular pressure. The contralateral forelimb was instmmented in 2 horses, and regional blood flow was determined at three consecutive times, with a minimum interval of 10 minutes between each determination (Group 2, Trials

1,2, 3), but without increasing intraarticular pressure. Cardiac output was determined prior to each experiment and mean arterial blood pressure was recorded before each trial. Horses were euthanized with an overdose of pentobarbital at the end of each study.

Instrumentation- Each horse was instrumented for drug administration and measurement of cardiac output by a thermodilution technique before induction of anesthesia 23. Briefly, a 7-

F, double-lumen, I lO-cm long thermodilution catheter^ was inserted through an introducer into the right jugular vein until the distal port was positioned in the pulmonary artery.

Polyethylene tubing (PE-240) for injection of cold solution was advanced through an introducer inserted in the right jugular vein proximal to the thermodilution catheter until the tip was positioned in the right atrium. Catheter position was verified by observing characteristic pressure waveforms. Horses were sedated with xylazine (0.5 mg/kg,IV) and anesthetized with 5% guaifenesin (IV, to effect) and thiamylal (4 mg/kg, IV) and maintained with sodiurn pentobarbital (5-15 mg kg-1 hr-1, IV). Horses were intubated, positioned in dorsal recumbency, and mechanically ventilated with 1(K)% oxygen. A 20- gauge catheter was inserted in a facial artery after anesthesia induction, for direct measurement of arterial blood pressure and collection of arterial blood samples for determination of arterial pH and blood gas^. The zero-pressure reference point for all pressure measurements was the point of the shoulder 24. Systolic, mean and diastolic arterial blood pressures were continuously displayed and arterial blood gases were

76 determined hourly. Mean arterial blood pressure was electronically derived. Cardiac outputs were determined hourly and before each experiment and expressed in ml-kg-i-min-i.

Measurement o f joint pressures Midcarpal joint pressure was measured by inserting a

20-gauge needle in the dorsal compartment of the midcarpal joint, 1 cm medial to the extensor carpi radialis tendon. We have documented that intraarticular pressures are equivalent in the medial or lateral compartment of the dorsal midcarpal joint pouch in h orses22. The needle was connected to tubing filled with heparinized 0.9% saline (heparin

10 lU/ml). Care was taken to ensure that the tubing was closed to the horse prior to insertion of the needle, in order to measure negative intraarticular pressure. The tubing was filled with heparinized saline to prevent clotting that could occur as a result of hemorrhage during needle insertion and was connected to a horizontally positioned pressure transducers placed level with the midcarpal joint (approximately 40 cm above heart level). The recorded pressures were continuously displayed on a physiograph^ precalibrated by a Hg column to allow measurements ranging from -10 to +80 mmHg pressures. Intraarticular pressure was increased by infusion of synovial fluid using a 20-gauge needle placed through the extensor carpi radialis tendon and into the midcarpal joint. Synovial fluid was collected from the tarsocrural joints of the same experimental horse immediately before use to avoid potential effect of temperature. Intraarticular pressures was increased and maintained at the predetermined values by continuous monitoring on the physiograph.

Measurement o f regional synovial membrane blood flow -Regional synovial membrane blood flow was determined using 15 [im-diameter polystyrene colored microspherese.

Red, blue and yellow were the selected colors. The median artery served as the injection

77 portal and the medial palmar artery as the arterial blood reference sample withdrawal portal.

The accessory cephalic vein was used to obtain venous blood reference samples for detection of arteriovenous shunts (Fig 1). The median artery was cannulated in a proximo-distal direction with a heparinized 22-gauge polytetrafluoroethylene catheter^. A second heparinized 22-gauge polytetrafluoroethylene catheter was inserted in a disto-proximal direction in the medial palmar artery. A third heparinized 22-gauge polytetrafluoroethylene catheter was also placed in the accessory cephalic vein for withdrawal of the reference venous sample (Fig 1).

Colored microspheres (red, blue or yellow) were thoroughly dispersed by vortex mixing for 30 seconds, suspended in 5 ml of saline containing 0.02% polyoxyethylene sorbitan monooleate (Tween 80) and separately injected at a constant rate in 2-3 sec into the inflow median artery catheter. Reference arterial blood samples were withdrawn for 60 seconds with a calibrated withdrawal pumps set at a withdrawal rate of 10.89 ml/min starting 10 seconds before microsphere injection. This rate was selected because it was much smaller than the flow to the limb (previously determined, approximately l(X) ml/min) and allowed collection of approximately 10 ml of blood in one minute, which simplified analysis according to the manufacturer’s recommendations*». The withdrawal rate of the pump was verified independently at the end of each experiment To ensure a desired microsphere count of 1000 spheres per tissue sample, 3x10^ (trial 1) and 6x10^ (trials 2 and 3) microspheres were used for injection. These values were estimated based on the percent weight of an average synovial tissue specimen of 2 g, compared to the average weight of the limb distal to the arterial injection portal of 60(X) g (0.033%), from which the total amount of microspheres to inject was calculated so that each tissue sample would contain approximately 1000 spheres (Number of microspheres to inject = 1000 x 100/

0.033= 3 X 106)*».

78 Collection of tissue - The joint capsule (including fibrous layer and synovial membrane) was collected inunediately after euthanasia from the dorsal, lateral and medial palmar midcarpal joint pouches, the dorsal antebrachiocarpal joint, and the dorsal rnd palmar metacarpophalangeal joint pouches. The synovial membrane (synovium and subsynovium) was dissected from the fibrous joint capsule and separated into 0.5-2.0 g specimens. One or two specimens were obtained from each site, discarding specimens that had evidence of irritation or bruising at the site of needle insertion. All sampled joints were inspected for evidence of cartilage fibrillation, or synovial membrane thickening to determine if inclusion criteria were met (see above).

Tissue processing - Reference blood samples and tissue specimens were digested and filtered for recovery of microspheres. Tissue digestion was performed as follows: after weighing, tissues specimens were placed in glass tubes and 7 ml of 4M KOH with 2%

Tween 80 were added to each tube. Blood samples were digested with 4 ml of 2% Tween

80,4.4 ml of 16M KOH and 2 ml of 20% Tween 80 for each 10 ml of blood. Blood and tissue specimens were allowed to digest overnight in a 3?o C water bath. Each digest was then filtered under vacuum using a 8-p.m 25-mm diameter polyester filter^ placed in a stainless steel syringe holder. Digestion tubes were rinsed with 2% Tween 80 and 70% ethanol to ensure collection of all spheres. The filter containing the microspheres was placed in a 1.5 ml Eppendorf tube, to which 100 pJ of NN-dimethylformamide was added to release the dye. Each tube was vortex mixed for 30 s and centrifuged for 3 minutes at

2000 g.

Microsphere assay technique and calculations- The concentration of each dye solution

(proportional to number of microspheres) was measured by spectrophotometry: at

79 wavelengths of 448 nm (yellow), 530 nm (red) and 672 nm (blue) using dimethyiformamide as the blank. Nine standards were used to develop the standard matrix before each assay. A matrix inversion technique was used to correct for overlap of composite spectra 25. Samples were diluted when necessary to maintain absorbance of each wavelength under 1.3 absorbance units (AU), to ensure linearity of the absorption-concentration curve according to the Lambert-Beer law 26. Ail tissue samples containing less than 4(X) spheres were discarded from analysis.

Blood flow to each tissue specimen was calculated as follows:

Blood flow to tissue sample (|i/min)=

AU of tissue sample x reference blood flowfml/min ) £1)

AU of reference sample.

Blood flow was expressed in pi min-i g-i of tissue lo.

Statistical analysis- Distribution of data for all sampled populations were tested for normal distribution using the Kolmogorov-Smimoff test for continuous data, and for equality of variances using the F test. The assumption of normal distribution was met in all instances.

However, the variances about the mean of midcarpal blood flow for trials 1,2 and 3 were unequal, with trial 1 showing a higher variability. Therefore a nonparametric test (Friedman test for repeated measures) was used to test for differences among trials. If a significant effect was detected, a Dunn’s comparison test was used to compare individual means 27.

Mean dorsal and palmar metacarpophalangeal pouch blood flow and mean fibrous capsule and synovial membrane dorsal midcarpal joint blood flow were compared using a two-tailed paired Student’s t test. The coefficient of repeatability of blood flow determination within each animal and among colors was determined from the control

80 animals using the mean square error of the analysis of variance table performed on the

log-transformed data as there was an association between the magnitude of measurement

and repeatability. A 95% confidence interval for the coefficient of repeatability was also constructed for each pair of color, using log-transformed data 28. The coefficient of

variation of blood flow within each animal was calculated for each site that had two tissue

specimens available for blood flow determination. The coefficient of variation among

animals was calculated from the pooled mean and standard deviation for each site. Mean

arterial blood pressure obtained before each trial was compared using a one-way analysis of

variance with repeated measures. Data are reported as mean ± SE. The level of significance

for all tests was set at P < 0.05.

Results

Cardiac outputs determined before the start of each experiment measured 51.7 ± 3.7

ml-min-i-kg-i. Mean arterial blood pressures recorded before each trial were 119.2 ± 3.0

(trial 1), 116.8 ± 3.7 (trial 2), and 116.5 ± 3.9 (trial 3). Mean PaCOi was 38.00 ± 1.57

mmHg and mean arterial pH was 7.41 ± 0.02 for all horses and all trials. There was no

significant difference in mean arterial blood pressures, pH and PaC02 among trials.

Regional synovial membrane blood flow- Baseline regional synovial membrane blood

flow for each joint was 103 ± 8 (dorsal radiocarpal), 108 ± 36 (dorsal midcarpal), 61 ± 12

and 50 ± 11 (lateral and medial palmar midcarpal joints) and 17 ± 3 and 26 ± 5 (dorsal and

palmar metacarpophalangeal joints) |il-min-i-g-i (Table 1). Blood flow to the dorsal aspect

of the metacarpophalangeal joint was significantly less than that of the palmar aspect

(p=0.048). Synovial membrane blood flow was significantly higher than fibrous capsule

flow in the dorsal pouch of the midcarpal joint (96 ± 32 vs 24 ± 6 |il-min-i-g-i, p=0.034)

81 (Fig. 4.2). Microspheres were not detected in venous blood reference samples obtained from the accessory cephalic vein at baseline or after increased intraarticular pressure.

Midcarpal joint pressure- Baseline intraarticular midcarpal joint pressures measured -2.86 ±

1.45 mmHg, with a median value of -2.00 mmHg and 95% confidence intervals of -6.42 and 0.70 mmHg. Increases in intraarticular pressures of 30 and 60 mmHg in the midcarpal joint resulted in a significant decrease (p=0.0224) in synovial membrane blood flow to

16% of baseline values (baseline: 108 ±36 |j.l-min-i-g-i, 30 mmHg: 12 ± 7 ml-min-i-g-i,

60 mmHg: 11 ±3 p.l-min-1 g-i)(Fig. 2). There was no difference between synovial membrane blood flow measured at intraarticular pressures of 30 and 60 mmHg. Sequential synovial blood flow of control limbs were not significantly different between trials (trial 1:

109 ± 5 1 pl-min-i-g-i, trial 2: 94 ± 48 |il-min-i-g-i, trial 3: 128 ± 52 pl-min-i-g-i).

Repeatability: The overall coefficient of variability for the method, calculated from log-transformed data from the group 2 horses on 39 samples from all sites was 47%. The

95% confidence interval for blood flow determination between the first and second blood flow determination was 13% and 27% (i.e. the blood flow could be expected to range between 13% below and 27% above the value obtained), between the second and third blood flow determination was 16% and 30% and between the first and 3rd determination was 21% and 22%.

Limits o f agreement for the same site: For 26 samples where 2 tissue specimens from the same site were available, the limits of agreement were determined from the log-transformed data, with 25 degrees of freedom, giving an interval of 8% below to 16% above the value obtained.

82 Coefficient o f variation- Within each horse, the coefficient of variation of blood flow for all sites averaged 29% (range 25% -34%). The mean coefficient of variation of blood flow between horse at each site was 64% (range 54%-73%).

Discussion

Our study is the first to examine synovial blood flow and tissue specific blood flow in joints with the use of colored microspheres. Regional synovial membrane blood flows in equine joints at baseline ranged from 17± 3 (dorsal fetlock) to 108± 36 (dorsal midcarpal joint pouch) p.l-min-* g-i (mean ± SE). These results are comparable to stifle and hip joint blood flows obtained in dogs as measured with radioactive microspheres 29-31.

Our synovial membrane tissue samples included synovium and subsynovium.

Previous studies have shown that capillary density is highest within 25 |xm of the joint boundary, and decreases thereafter 32,33. Our tissue samples were obtained by gross dissection, and thus contained synovium and subsynovium. The inclusion of less vascular tissue may have resulted in lower blood flow values than if only the most superficial, capillary dense, 25 |im of synovial membrane had been obtained.

We were able to demonstrate significant heterogeneity of blood flow from different sites within a joint. Blood flow to the dorsal aspect of the metacarpophalangeal joint was significantly lower than that of the palmar aspect Morphologically, the synovial membrane of the dorsal aspect of the metacarpophalangeal joint is classified as fibrous, because the synovium rests on a fibrous, relatively avascular subsynovial layer 2. In contrast, synovium of the palmar aspect of the fetlock joint pouch is classified as areolar as it contains numerous synovial villi resting on areolar subsynovial tissue. Synovial villi are very vascular, which correlates well with our findings of a higher blood flow at that site.

Our study further characterized the blood flow to specific tissues within the midcarpal joint.

83 and demonstrated that blood flow to the fibrous joint capsule is lower than that of the synovial membrane.

An intraarticular pressure of 30 mmHg was sufficient to cause a significant and marked reduction in articular tissue blood flow in horses in dorsal recumbency. The effect of hydrostatic pressure, capillary pressure and thus blood flow occlusion pressure may vary with limb and body position. Venous pressure in the feet of human beings standing absolutely still approximates 90 mmHg,34 and it can be inferred that a similar or greater value would be obtained in standing still horses. Locomotor activity decreases venous pressure in the feet of adult humans to less than 25 mmHg because of the combined effects of the pumping action of skeletal muscles and venous valves 34. We are confident that a similar effect would occur in active horses. We have measured the capillary pressure at the level of the metacarpophalangeal joint in laterally recumbent horses by the venous occlusion technique, and obtained values of 13.3 mmHg 35. In another study, the more rigid tissues of the distal digit (hoof) of laterally recumbent horses yielded capillary pressures of 37 ± 2 mmHg. This capillary pressure is higher than that in the dog hindpaw, and was counterbalanced by a higher tissue pressure of 25.6 ± 2.5 mmHg, thus preventing edema formation 36. In our study, the use of an upright limb position may have resulted in a progressive increase in capillary hydrostatic pressure because of inactivity, thus altering the intraarticular pressure necessary to occlude blood flow.

The formation of synovial fluid, an ultrafiltrate of plasma, is dependent on synovial capillary pressure as there is no synovial basement membrane L Capillary pressure in our study was likely less than 30 mmHg, such that reduction of blood flow was significant with intraarticular pressures of 30 mmHg or more. We speculate that a reduction in blood flow may lead to decreased capillary hydrostatic pressure, decreased filtration and consequently decreased synovial fluid production. In addition, articular clearance of small

84 solutes which are dependent on venous flow would be expected to be decreased. At intraarticular pressures of 25 cm H 2O, morphologic synovial capillary compression has been observed 37. The deleterious effects of increased intraarticular pressure on synovial fluid metabolite accumulation has been reported Relative joint ischemia resulted in synovial fluid acidosis, increased CO 2 pressure, and lactate accumulation, reflecting poor clearance and anaerobic metabolism. Although it was initially reported that intraarticular pressures above systolic blood pressure were necessary to obtain these effects, subsequent studies have shown that intraarticular pressure greater than 45 mmHg resulted in significant lactate and metabolite accumulation lA These previous studies served as the basis for our choice of intraarticular pressures above and below 45 mmHg. Geborek 38 reported a significant decrease in synovial blood flow (measured by doppler flowmetry) when intraarticular pressure was increased to 20 mmHg. Our results corroborate that relatively low intraarticular pressures can produce significant reduction in blood flow. The effect of intraarticular pressure on synovial membrane blood flow is, however, dependent on synovial membrane capillary hydrostatic pressure, which in turn will vary with positioning, locomotor activity, vasomotor tone and venous pressure. Our findings suggest that articular decompression and resolution of joint effusion may be beneficial as part of the treatment of joint disease.

There was marked variability in individual blood flow values between horses in the present study. This finding is similar to that of studies performed with radioactive microspheres, where blood flows from the same sites varied between animals, and was suggested to be function of animal variation rather than variability in the method 39-41.

Differences in vascular tone or cardiac output may contribute to these differences. Blood flow values were more consistent within each horse and suggest that colored microspheres

85 are a useful method for measurement of synovial membrane blood flow in the horse when a baseline value is obtained.

Microspheres have been used successfully to measure regional blood flow to several organs, including bone, myocardium, endocardium, lung, ocular and renal tissues

12,41-46. High agreement has been demonstrated between blood flow measured by radioactive and colored microspheres 12. Colored microspheres eliminate the need to use radionuclides, use different colors to allow serial measurements, and are biocompatible, such that chronic implantation is possible. Other methods of articular blood flow determination include venous occlusion plethysmography, clearance of rapidly-diffusing radioactive solute (23Na, i3ii,i32Xe, tritiated water) or heat, and laser Doppler flowmetry

8.38. These methods provide relative estimates of perfusion, assume homogeneity of tissue, do not account for flow to skin or periarticular tissues (articular soft tissues are estimated to form only 15% of an articular segment 47.48), and in the case of clearance studies, assume that solute clearance is only flow limited. The use of microspheres for synovial blood flow measurement allows an absolute measure of blood flow, which can be determined from small tissue samples, and allows quantification of flow from different anatomical sites or within different tissue types.

Aggregation of microspheres, maldistribution of blood flow including lamination of flow, and arteriovenous shunting are problems that may be encountered when using a microsphere method to determine blood flow. Aggregation of spheres results in nonuniform distribution of the microspheres and may occlude blood flow to the tissue of interest. Microsphere aggregation is prevented by the addition of the surface-active polyoxyethylene sorbitan monooleate (Tween 80). The cardiovascular effects of Tween 80 are virtually non-existent at the concentration used in our study (0.02%), particularly since regional rather than systemic injection was performed 49. Cardiac output and mean arterial

86 blood pressure values throughout our study were comparable to those obtained in other studies in horses under general anesthesia 5041. We believe that maldistribution of blood flow, which may be caused by the laminar flow of microspheres within the inflow artery, was insignificant, because the coefficient of variation of blood flow was small within each horse for a given site. We were unable to demonstrate arteriovenous shunting in any horse in our study during altered blood flow conditions such as increased intraarticular pressure.

In conclusion, this study successfully used colored microspheres for the measurement of articular blood flow in horses. Significant heterogeneity of blood flow distribution was demonstrated between the synovial membrane and fibrous joint capsule.

Increased intraarticular pressure to 30 mmHg maricedly decreased synovial membrane blood flow. Increased intraarticular pressure may contribute to synovial membrane pathology via an ischemic pathway.

Footnotes a Swan-Ganz thermodilution catheter, Columbus Instruments, Columbus, OH b ABL 500, Radiometer Copenhagen, Copenhagen, Denmark c Spectramed P23XL Transducer, Oxnard, CA d Grass Model 7D Polygraph, Grass Instmments Co, Quincy, Mass e Dye-Trak microspheres, Triton Technology Inc., San Diego, CA f Angiocath, Becton-Dickinson, Sandy, UT g Model 940 Infusion/Withdrawal pump. Harvard Apparatus Co, Dover, Mass h Triton Technology Inc., San Diego, CA i Beckman D-70 Spectrophotometer, Scientific Instrument Division, Beckman Instrument, Fullerton, CA

87 Figure 4.1: Medial aspect of the left forelimb of a horse illustrating normal anatomy and instrumentation for microsphere determination. The scale indicates approximate height above the heart of the metacarpophalangeal joint.

88 64 cm

Carpus Median artery Antebrachiocarpal joint Midcarpal joint

Medial palmar artery

Forearm Metacarpo phalangeal joint Reference syringe

89 Figure 4.2: Synovial membrane blood flow in the dorsal pouch of normal equine midcarpal joints in dorsal recumbency at a joint angle of 135° after increased intraarticular pressure (lAP) (Group 1) or in normal joints (Group 2). Data are mean +/-SE.

* indicates significant difference from baseline

90 0.20 T

0.18-

0.16- A Control ï 0.14- M Q Increased lAP a 0.12- I H I 0-10"

I 0.08- z 0.06- IAP=tiO mmHg

0.04- lAP = 30 mm Hg 0.02 - □ 0. 0 0 - ' I 1----- Trial 1 Trial 2 Trial 3

91 Figure 4.3. Scattergram of the coefficient of variability between three simultaneous blood flow determinations obtained with three different microsphere colors, plotted against number of microspheres. This graph illustrates the higher variability associated with low microsphere numbers.

92 25 n

■o 2 0 -

15-

1 0 -

5 -

500 1000 1500 2000 2500 Microsphere number

93 Figure 4.4. Bland and Altman plot of the difference between three serial (one minute interval) blood flow determinations in the same tissue specimen, to illustrate the repeatability of blood flow determinations. Indicated are the mean difference and 95% confidence interval for serial blood flow determinations.

94 Difference (pi min o Ù) o in o f Z o &

O U l Joint Tissue n Blood flo w lil/m in/g Dorsal radiocarpal Syn 16 103 ± 8 Dorsal midcarpal Syn 8 108 ± 36 Dorsal midcarpal Fibrous 8 24 ±6* Lateral palmar midcarpal Syn 7 61 ± 12 Medial palmar midcarpal Syn 9 50± 11 Dorsal metacarpophalangeal Syn 15 17 ±3** Palmar metacarpophalangeal Syn 15 26 ± 5

Table 4.1 Blood flow to articular soft tissues of normal midcarpal and metacarpophalangeal joints in horses positioned in dorsal recumbency, with the midcarpal joint at 135° angle. Data are mean ± SE.

Syn = synovial membrane (synovium and subsynovium). Fibrous = fibrous joint capsule.

* Midcarpal joint fibrous capsule blood flow significantly lower than midcarpal joint synovial membrane

(P = 0.034)

** Dorsal metacarpophalangeal joint blood flow significantly lower than palmar (P = 0.048) References

I. Ghadially, F. N. Structure and function of articular cartilage. Clin Rheum Dis 7:3-27, 1981.

2. Hasselbacher, P. Structure of the synovial membrane. Clin Rheum Dis 7:57-69, 1981.

3. Lund-Olsen, K. Oxygen tension in synovial fluid. Arthritis Rheum 13:769-776, 1970.

4. Levick, J. R. Hypoxia and acidosis in chronic inflammatory arthritis; relation to vascular supply and dynamic effusion pressure. J Rheumatol 17:579-582, 1990.

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100 CHAPTER 5

LOCAL HEMODYNAMICS, PERMEABILITY AND O2 METABOLISM OF INNERVATED OR DENERVATED ISOLATED JOINTS IN AN IL-I MODEL

Introduction

Synovial joints form functional units that facilitate relative motion of masses under load. I The blood flow, fluid exchange and innervation of joints have adapted to this biomechanical role and to the preservation of a supply of nutrients to a tissue under compressive load, the articular cartilage. Articular cartilage is an avascular, aneural but metabolically active tissue. Nutrients to cartilage must cross the blood-joint barrier, which consists of a fenestrated capillary endothelium in series with the overlying synovial intima, and in parallel with the synovium interstitium. In turn, the exchange characteristics of the blood-joint barrier can be modified by alterations in plasma and synovial fluid composition, capillary and joint cavity hydrostatic pressure and capillary and synovial membrane permeability. The means of acutely affecting these changes are through blood flow and innervation.

A Starling’s based mechanism has been proposed and substantiated to explain the pathways of fluid exchange across the blood-joint barrier. Starling proposed that the rate of fluid exchange across a physiologic barrier is the result of differences of hydrostatic and oncotic pressures acting across a semi-permeable barrier and modified by the filtration and oncotic reflection coefficients of the tissues. The influence of various osmotic fluids on joint fluid filtration and intraarticular pressure have clearly supported the use of Starling’s forces to explain joint fluid exchanges. The presence of local heat, periarticular edema and

101 joint effusion in articular disease indicates an imbalance between vascular permeability and tissue clearance that leads to tissue swelling, which warrants further investigation.^

Separate clinical and experimental studies have examined the physiology of blood flow, innervation, and fluid exchanges of joints in health and their alteration with disease.3-

6 However, these mechanisms have not been examined in an isolated model or during acute articular inflammation. Extracorporeal isolated organ preparations improved our understanding of co-dependent physiopathologic mechanisms of organs such as heart,? lung, liver, intestines and others.

In this study, we use an isolated joint organ preparation to examine pathophysiologic alterations of the joint, including blood flow, permeability, fluid exchanges, and oxygen metabolism associated with innervation or denervation in a model of acute joint inflammation.

Materials and Methods

H orses -Twenty-four adult horses (body weight 330-550 kg, 8 females, 8 geldings, 8 stallions) of various breeds were used. Horses were determined free of metacarpophalangeal joint disease by thorough clinical examination, synovial fluid analysis, and gross post-mortem examination. All experiments were conducted in accordance with the Animal Care and Use Guidelines of The Ohio State University.

Experimental procedures

Horse instrumentation -Horses were instrumented for cardiac output measurements and intravenous drug administration prior to induction of general anesthesia. Horses were sedated with xylazine hydrochloride (0.25 mg/kg) and the skin over both jugular furrows

102 was clipped, aseptically prepared, and catheter insertion sites were locally infiltrated with lidocaine hydrochloride. A flow-directed thermodilution catheter was inserted in one jugular vein through a catheter introducer and advanced until the distal tip was located in the pulmonary artery. This catheter was used for thermal dilution estimates of cardiac output.9

Polyethylene tubing for administration of cold dextrose solution was inserted proximal to the thermodilution catheter and advanced until the distal tip was positioned in the right atrium. Catheter placement was guided and position was confirmed by observation of characteristic pressure waveforms. The opposite jugular vein was cannulated with a 14 gauge teflon catheter for drug administration. Cardiac output was recorded as the mean of 3 measurements, using a cardiac output computer». After induction of general anesthesia, a facial artery was cannulated for direct measurement of arterial blood pressure.

Anesthesia and monitoring

Only one horse was studied on a given day. Horses were sedated with xylazine

(0.5 mg/kg) and anesthesia was induced with guaifenesin (25 mg/kg) followed by sodium thiopental (2 mg/kg, iv). Anesthesia was maintained with sodium pentobarbital (loading doses 4 mg/kg, maintenance 5-15 mg/kg/hour, iv, to effect) and controlled ventilation with

100% oxygen. A bipolar ECG (base-apex lead) was recorded throughout each study for determination of heart rate and rhythm. Arterial pressure and ECG were continuously displayed and semicontinuously recorded on a multichannel recorder^. Systemic arterial blood gases, packed cell volume and total protein were determined hourly. Intravenous poly ionic isotonic fluids were administered (5-10 ml/kg/hr) throughout each study. Mean systemic arterial pressure was maintained at > 70 mm Hg, and ventilation was adjusted to maintain PCO2 < 55 mmHg. Heparin (50,000 lU iv) was administered every 90 minutes.

103 Joint isolation procedure and instrumentation (Fig.5.1)

One metacarpophalangeal joint (MCP) was randomly selected and the horse was positioned with the selected limb lowermost. All circuit tubing was thoroughly flushed with heparinized saline. The medial palmar artery and vein were isolated at the level of the proximal third of the metacarpus. The artery was caimulated with polyethylene tubing (PE

320) in a disto-proximal direction proximal to the anticipated site of joint isolation, and in a proximo-distal direction distal to the anticipated site of joint isolation. Blood was diverted from the cannulated artery, circulated through a precalibrated peristaltic pumpc and infused via the distal cannulated medial artery to the joint preparation. In a similar fashion, the medial palmar vein and the medial and lateral palmar digital veins (PE 240) were cannulated and connected to a Y-connector and venous outflow was diverted to a reservoir and reinfused to the horse via the median vein using a second peristaltic pump. Venous pressure was controlled by adjusting the height of the extracorporeal reservoir. The medial palmar digital artery was cannulated for collection of a reference sample for blood flow determination by the microsphere method (see below). Final isolation of the joint was achieved by transection of all soft tissue, sectioning of the bones at the level of the midmetacarpus and dislocation of the proximal interphalangeal joint. In innervated preparations, the medial and lateral palmar nerves were isolated and preserved.

Hemorrhage was controlled by electrocoagulation or ligation. A 20 gauge 2.5 cm needle was inserted in the MCP joint using a palmaro-medial approach by insertion of the needle between the distal aspect of the medial sesamoid bone and the palmar aspect of the first phalanx, immediately distal to the medial metacarpo-sesamoidean ligament After baseline synovial fluid collection, the needle was connected to a heparinized saline filled tubing connected to a three way stopcock and pressure transducer, for continuous intraarticular measurement (first 240 minutes) and for synovial fluid collection (at 240,300 and 330 minutes). The isolated joint was suspended from a calibrated FT03 transducer connected to

104 a physiographd for continuous measurement of weight changes. All circuit pressure transducers were placed at the level of the joint preparation. Initial circuit arterial flow was set at 10 ml/min until completion of the preparation and establishment of isogravimetric state (see below). This joint isolation technique established a pressure-driven auto-oxygenated, pump-perfiised, extracorporeal isolated joint model.

Experimental protocol

The joint preparation model was randomly assigned to one of four groups:

1-Control; 2) Control-denervated; 3)Inflamed; 4)Inflamed-denervated. Inflammation was induced by intraarticular injection of 0.35 ng/kg BW of recombinant human interleukin-1 Be

(11-16). After joint isolation, isogravimetric state (no gain or loss of weight in the isolated preparation) was established and maintained for 20 minutes by adjustment of arterial flow. 10 The preparation was maintained for 6 hours, after which horses were euthanized with an overdose of pentobarbital (100 mg/kg, iv) and specimens collected. Baseline (time

0) was considered as the time after establishment of isogravimetric state and collection of baseline synovial fluid sample.

Sample collection

Circuit arterial and venous blood samples were collected at baseline, and hourly thereafter, in heparinized syringes, and transported on ice for blood gas determination.

Packed cell volume and total protein (refractometer) were measured in the circuit arterial blood. Synovial fluid samples were collected after joint isolation at baseline (time 0), 240,

300 and 330 minutes into appropriate vials as described below. All available fluid was collected at each collection time. Between the beginning of the experiment (time 0) and 240 minutes, synovial fluid was allowed to accumulate in the joint, and intraarticular pressure

105 was continuously recorded, using a pressure transducer placed at the level of the isolated joint and connected to a precalibrated physiograph module^. Local hemodynamic and

oxygen metabolism measurements were obtained hourly. Synovial membrane blood flow

was obtained at time 0,60 and 330 minutes. Synovial membrane permeability to albumin

and dextran was determined from 330 to 360 minutes. At 360 minutes, the preparation was

disassembled and weighed, and each joint was carefully examined for evidence of

abnormalities such as cartilage lesions, synovial membrane proliferation, and periarticular

osteophyte formation. The presence of such findings excluded the horse from the study.

Synovial membrane specimens were obtained for fluorescence studies and determination of

wet/dry ratio. Specifics of each sample and specimen collection and analysis are described

below.

Local hemodynamics measurements and calculations

After baseline measurements were recorded, circuit arterial pressures and flows

were recorded every 5 minutes for one hour to examine the influence of n IB on local

articular hemodynamics. Circuit arterial and venous pressures were then adjusted as needed

to maintain pressures at isogravimetric state, respectively, and circuit arterial flow values

were recorded hourly to determine alterations in hemodynamics. Total vascular resistance

(TVR) of the isolated joint was calculated as follows: TVR (mm Hg/ml/min) = (Pan - Pven V Q art (I) where Pan is the arterial pressure to the circuit (mm Hg), Pven is the venous pressure and

Q art is the arterial flow to the circuit (ml/min).

Synovial fluid collection was re-initiated at 240 minutes, and production of

synovial fluid was measured from 240 to 300 minutes, and 300 to 330 minutes by

collection into a graduated cylinder.

106 Synovial fluid total protein were measured using the BCA methodC Synovial membrane permeability surface area product (PS)i was estimated at 330 minutes using the following calculation:

- ^ = PS (Cp - Cf) (2) where is the protein flux across the synovial membrane, and Cf and Cp are the total protein concentration in synovial fluid and arterial plasma, respectively.

The protein flux was measured by the following equation:

^ = V * d C # (3) where V is the volume of synovial fluid produced between 300 and 330 minutes, and dCf is the difference in synovial fluid protein concentration between 300 and 330 minutes. This time period was chosen because lAP was 0 mm Hg (atmospheric presstue) during that time and synovial fluid flow was not influenced by the increased LAP observed at 240 minutes.

Oxygen metabolism measurements and calculations

Circuit arterial and venous blood were analyzed for PO 2, PCO2 and pH and corrected to body temperature at the time of collection. Total hemoglobin (Hb) and arterial oxygen saturation (Sa 0 2 ) were measured in each sample using a co-oximeterg, and arterial oxygen content (Ca 0 2 ) in each sample was calculated as follows:

Ca02 (ml/dl)= Hb x Sa02 x 1.36 + PaÛ 2 x 0.003. (4)

Circuit oxygen delivery (DO 2) was calculated as

DO2 (ml/min/g)= Ca02 (ml/dl) x Qan (ml/min)/ W (5) where Qan is the arterial flow and W is the weight in grams of the isolated joint. Circuit oxygen consumption (VO 2) was calculated as

107 VÛ2 (ml/min/g) = C(a-v) O 2 x Q art /W, (6) where C(a-v) O2 is the arteriovenous O 2 content difference. Oxygen extraction ratio (O 2

ER) was calculated as

O2 ER= C(a-v) 02/Ca02. (7)

Synovial fluid blood gases were measured at baseline, 240,300 and 330 minutes.

Synovial fluid O 2 gradient was calculated as:

O 2 gradientsF= Pa02 - PSO 2 (8 ) w here PsÜ2 is the oxygen tension in synovial fluid at each time period.

Determination of bloodflow by colored microspheres

Blood flow to synovial tissue was determined at baseline after joint isolation, 240 and 330 minutes, using a colored microsphere technique.^ Blue, red or yellow microspheres were injected into the circuit arterial port and a reference blood sample was simultaneously collected from the medial digital artery reference sample port Synovial tissue collected at the end of the experiment (15 nun x 10 rrun) was dissected and blood flow to this sample was determined. Briefly, blood and synovial tissue were digested KOH and ethanol, filtered on 8 jim polystyrene filter. Dye was then released from the microspheres using dimethylmethylformamide, and the concentration of the dye, proportional to the concentration of microspheres, was determined by spectrophotometry, after standardization and correction for overlap of individual spectra. Blood flow was then calculated as follows:

Blood flow to = AU of tissue sample X reference blood flow (ml/mint (8) tissue sample (p.l/min) AU of reference sample

Blood flow was expressed in pJ/min/g of tissue.

108 Synovial membrane permeability to FITC-dextran (M W 144,(XX))

Synovial membrane permeability to dextran (MW 144,000) was measured by injection of 2 ml of a 0.5 mg/ml FTTC-Dextran solution in normal saline, prepared before each experiment, and kept protected from light in an amber bottle. A circuit venous blood sample was obtained immediately before injection, every 2 minutes for 20 minutes, and every 5 minutes for another 10 minutes after injection. All vials were protected from light, and serum immediately separated, stored in covered vials and frozen until assay.

FTTC-dextran concentrations were measured by spectrophotofluorometryj with excitation/emission wavelengths of 485 nm and 530 nm respectively. All background values were subtracted for analysis. Normal horse serum was used to prepare standards.

Synovial membrane permeability to albumin

Synovial membrane permeability to albumin was determined by intraarticular injection of 2 ml of Evans blue-albumin (40 mg/ml of bovine serum albumin, MW

68,(XX)), performed at 330 minutes, and allowed to absorb for 30 minutes. Evans blue-albumin was prepared by dissolving 0.6 g of Evans blue and 4 g of bovine serum albumin in 0.9% NaCl solution, a solution which results in negligible amounts of free dye. 11 After euthanasia, sections of synovial membrane and underlying fibrous capsule (20 mm X 10 mm) were harvested from the dorso-medial aspect of the MCP joint, packed in

OCT and snap frozen in liquid nitrogen and preserved at -70C. Frozen sections (7pm) were prepared on poly-L-Lysine coated slides using a sliding microtome.

Amount and depth of Evans blue fluorescence of the synovial capsule (intima, subintima, and fibrous layer) was analyzed on a computer image acquired with a 5 watt argon laserh and image analysis system» at an excitation wavelength of 488 nm, and

109 collection of fluorescence with a 605 nm long pass filter. Scanning was performed at a microscope setting of 40x, with step sizes of 4jim, and an image size of 500 x 300 points.

On each specimen, background tissue fluorescence was measured, and depth of dye penetration was recorded as the distance from the synovial intima to a point in the subsynovial tissue where fluorescence was equivalent to background. Five measurements, separated by a 20 nm distance,were obtained in the villous and fibrous synovial membrane respectively of each specimen, and fluorescence was reported as the mean of these 5 measurements for each region.

Wet/dry ratio

Synovial specimens (20 mm x 10 mm) were harvested from the dorsal aspect of the

MCP joint, weighed and dried in an oven at 80° C for 48 hours and the ratio of wet to dry weight was calculated.

Statistical analysis

A three factor analysis of variance (model (irmervated or denervated), treatment

(control or inflamed) and time) with repeated measures (time or site) was used to test for differences among groups and across time in the means of systemic and circuit hemodynamics, systemic and circuit blood gas tensions, synovial fluid total protein, oxygen metabolism parameters, FTTC-dextran permeability and depth of fluorescence of

Evans blue albumin between sites. Assumptions of normality and homogeneity of variances were not met for synovial fluid production and for intraarticular pressure, so data were normalized for these parameters by log transformation (synovial fluid production) or square root transformation (intraarticular pressure) prior to analysis. A two factor analysis of variance (model, treatment) was used to test for differences in the means of synovial fluid production, intraarticular pressure at 4 hours, synovial membrane PS and wet-dry

110 ratio. For all parameters, if a significant F test was obtained, a post hoc Student Neuman

Keuls multiple comparison test was used to detect individual differences among means.

Significance for all tests was set at P<0.05, and trends are reported when P< 0.15. Data are expressed as mean ± SE.

Results

Systemic hemodynamics

Systemic mean arterial pressure and PaCOi were maintained within stated limits of mean arterial pressure >70 mmHg and PaC02 < 55 mmHg (Table 5.1). There were no difference detected in systemic hemodynamics or blood gas analysis measurements among groups.

Local dynamics and fluid exchanges

Isogravimetric state was established for all joints for 20 minutes before the beginning of each experiment Isogravimetric state was obtained at a circuit arterial flow of

14.29 ± 4.64 ml/min, circuit arterial pressure of 151.45 ± 2.42 mmHg, and circuit venous pressure of 10.12 ± 0.40 mmHg (Table 5.2). There was no difference among groups in baseline isogravimetric state.

From time 0 to 60 minutes (when circuit flow was not adjusted to control pressure), significant changes in arterial pressure or total vascular resistance were not detected over time. However there was a trend (P=0.12) for a progressive increase in arterial pressure in the inflamed innervated groups (groups 3) (Fig 5.2), and a trend for increased arterial (P=0.11) and total vascular resistance (P=0.10) in the innervated groups

(Groups 1 and 3) (Fig 5.2).

During the remainder of the study period, circuit arterial blood flow significantly increased over time in all groups, from 14.29 ± 4.64 to 27.24 ± 8.06 ml/min (Table 5.2).

Ill There was no difference in this parameter among groups. Similarly, total vascular resistance significantly decreased over time in all groups (P=0.009), and there was no difference among groups in this parameter (Fig 5.3). Blood flow to the synovial membrane was significantly increased (P=0.013) over baseline at 60 and 330 minutes in all groups (Fig 5.4). There was no difference in blood flow to the synovial membrane among groups.

Synovial fluid production was significantly increased in the inflamed groups

(groups 3 and 4) compared to the control groups (groups I and 2) (24.81 ± 7.7 ml vs 5.6

± .64 ml respectively, P=0.001), and there was no difference detected with denervation in this parameter (Fig 5.5). Intraarticular pressure was significantly greater at 240 minutes in the inflamed groups (groups 3 and 4) compared to control groups (groups 1 and 2) ( 6.46

± 1.65 mm Hg vs 1.21 ± 0.41 mmHg, respectively, P=0.005), and there was no difference detected with denervation (Table 5.3).

Synovial fluid total protein significantly increased over time (P<0.0001). There was no difference detected in synovial fluid total protein among groups (Table 5.3).

Synovial membrane permeability surface area product was significantly increased with inflammation (Groups 3 and 4 ) compared to control (Groups 1 and 2). Permeability surface area product was 0.00057 ± 0.0001 ml/min in control joints (Groups 1 and 2) and

0.0037 ± 0.00085 ml/min in inflamed joints (Groups 3 and 4) (P= 0.0002).

Parameters o f oxygen metabolism

Oxygen delivery (DO 2 ) significantly increased (P < 0.0001) in all groups over time

(Fig 4.5) and was not different among groups. Oxygen consumption (VO 2 ) significantly increased over time in all groups, however, there was a significantly greater increase

(P=0.01) in oxygen consumption in inflamed joints (groups 3 and 4) (Fig 5.7). There a trend (P=0.10) for increased oxygen consumption in innervated joints (groups 1 and 3).

112 Oxygen extraction ratio (0% ER) was significantly increased (P = 0.04) for the first three hours in inflamed joints (groups 3 and 4) (Fig. 5.7). There was a trend (P=0.09) for increased extraction ratio in denervated joints (groups 2 and 4).

Synovial fluid PaOi and synovial fluid pH were significantly decreased in all groups at 240 minutes (P<0.005) and was not different among groups (Table 5.3). There was a significant effect of model, treatment and time on synovial fluid 0% gradient. In control groups (Groups 1 and 2) 0% gradient significantly decreased at 240 and 300 minutes with denervation (P= 0.0(X)I). Inflammation significantly decreased synovial fluid

O2 gradient (P=0.01). In inflamed groups (Groups 3 and 4) O 2 gradient significantly increased with denervation (P=0.02).

Wet/dry ratio

Synovial tissue wet/dry ratio was significantly higher in the denervated group

(groups 2 and 4) (P=0.009) (Table 5.4). Denervated joints (Groups 2 and 4) had approximately 7% more edema fluid than innervated joints (Groups 1 and 3).

Permeability of the synovial membrane to albumin

The preferential pathway of albumin absorption was localized at the synovial villi.

Depth of penetration of Evans blue-albumin was significantly greater in inflamed joints

(groups 3 and 4) (P=0.023), and was most marked in the villous area (P=0.0003) (Figure

5.8 and Table 5.5).

FITC-dextran permeability

There was a significant increase in FTTC-Dextran concentration in the venous effluent of innervated joints (groups 1 and 3) (P=0.02). There was a trend (P=0.07) for

113 increased permeability in inflamed joints (groups 3 and 4) (Fig 5.9). Denervation decreased permeability to FTTC-dextran by approximately 26%.

Discussion

The principal findings of this study are a significant effect of denervation to decrease synovial membrane permeability to large molecular weight dextrans, and to induce synovial membrane edema. Inflammation increased parameters of oxygen metabolism, synovial fluid production, and synovial membrane permeability to albumin, and increased synovial membrane permeability surface area product of proteins.

Our findings of increased permeability to a large molecule such as dextran

(MW: 144,000) in innervated joints suggests a role of innervation in molecular selectivity of the blood-joint barrier.

The rate of appearance of dextran in venous blood is a function of the rate constant for fluid absorption from the joint and rate of removal from the interstitium of the synovial membrane into capillaries. Intraarticular hydrostatic pressure can significantly increase fluid absorption from the joint, and fluid absorption becomes a linear function at lAP greater than 9 mm Hg, a pressure that has been termed the breaking point or yield pressure. In the present study, lAP was returned to and maintained at atmospheric pressure after 240 minutes. It is possible that the increased lAP obtained at 240 minutes with inflammation caused the opening of interstitial channels that remained open after return to baseline.

However, the greater effect of innervation to increase permeability of large molecular weight dextran necessitates an alternative explanation. Synovial capillaries are richly innervated with sympathetic as well as small C-fibers.i3.i4 Experimentally induced plasma extravasation by intra-articular infusion of bradykinin is decreased by approximately 60%

114 with sympathetic denervation of the joint. I5.i6 Furthermore, bradykinin-induced plasma extravasation is dependent on intact postganglionic sympathetic fibers. *6 In our model, postganglionic fibers are intact in the joint capsule, such that the local vasoactive response mediated by postganglionic sympathetic fibers is preserved. In acute denervation, such as in that performed in this model, neuropeptides stored at nerve endings^? may still have been released in response to stimulation. Neuropeptides such as substance P and calcitonin gene related peptide (CGRP) are known to induce many of the changes occurring in acute inflammation, namely, vasodilation, increased vascular permeability, infiltration of leucocytes in venules, stimulation of phagocytosis, and mast cell degranulation.5.18

CGRP, but not substance P was shown to have a significant effect on articular vascular permeability. CGRP and substance P are collocated on small C-fiber nerve endings and can be released in the face of a stimulus such as interleukin-1. Chronic denervation by neonatal capsacin has no effect on experimentally induced plasma extravasation. Therefore sympathetic denervation provides an explanation of our findings of decreased

FTTC-dextran in denervated joints in venous outflow from denervated joints.

The influence of denervation to increase the tissue wet dry ratio, a measure of tissue edema, can also be explained by a decreased absorption of fluid from the interstitium. In our preparation, absorption of large molecules was decreased with denervation, which may also decrease convective flow of fluid, leading to tissue edema.

Intraarticular inflammation produced significant increases in oxygen metabolism of the joint unit. Oxygen delivery and extraction ratio provide an index of the efficiency with which tissues can extract oxygen for a given metabolic rate.i9 In normal tissues at normal or high levels of delivery, oxygen consumption is constant and independent of delivery. If delivery decreases or metabolic demand increases, tissue extraction ratio increases. In the present study, when oxygen delivery was below 0.61 ml/min/g significant increase in

115 extraction ratio was required to meet the increased oxygen demand of inflamed joints. As delivery increased, extraction ratio subsequently returned to baseline and still met O 2 demand (Fig 5.7). Although intraarticular pressure was allowed to increase during the first four hours of the study, this increase did not affect the ability of the tissue to increase extraction ratio in response to demand. Furthermore, the resulting lAP of 6.46 ± 1.65 mm

Hg observed at 4 hours in inflamed joints was well below the values of 20 mm Hg3 and 30 mmHg4 reported in previous studies to significantly decrease synovial membrane blood flow. However, in clinical situations, where equine metacarpophalangeal joint intraarticular pressure increases of greater than 30 mmHg have been reported,20 decreased synovial membrane blood flow and maldistribution of flow within the joint may result in a defect in extraction efficiency, leading to oxygen debt and anaerobic metabolism .21 In our study, the absence of a difference in synovial fluid oxygen tension in inflamed compared to control joints is explained by the low and insufficient increase in lAP to induce such changes .22 Other mechanisms by which extraction efficiency may be affected include a decrease in capillary density due to capsular fibrosis in chronic arthritis, edema formation resulting in increased diffusion distance, perivascular cuffing caused by severe inflammatory changes, microthrombi formation, or in bacterial arthritis the direct effect of

LPS to decrease cellular oxygen extraction efficiency. When extraction ratio increases in response to increased demand, a greater dependency on delivery exists. If demand continues to increase, for example if an inflamed joint is exercised, extraction ratio reaches a maximal value, and a critical level of oxygen delivery can be reached below which anaerobic metabolism is initiated, contributing to the pathology of the disease.

Synovial tissue blood flow values in our study in latter recumbency were greater than that previously reported for the equine dorsal MCP joint in dorsal recumbency.

Baseline blood flow in control-innervated joints were 43 ± 17 p.l/min/g whereas previously

116 reported values were 17 ± 3 pJ/min/g.4 In the study with horses in dorsal recumbency, the fetlock joint located approximately 64 cm above the heart base. In our study, isolated limbs where placed horizontally, but more importantly flow to the limb was controlled to maintain physiologic arterial pressures.

Synovial fluid is formed by ultrafiltration of plasma in the fenestrated capillaries of the synovial membrane. Increases in arterial pressure and to a lesser degree venous pressure significantly increase synovial fluid production's by increasing capillary hydrostatic pressure. In inflammatory processes, synovial fluid production is increased by increasing capillary permeability. In our study, the increased synovial fluid production was associated with an increase in articular pressure, however, as was previously reported, the relationship between increased volume and pressure is non-linear,24 such that predicting intraarticular volume from pressure is not possible unless that relationship is known.

The permeability surface area product of the blood joint barrier reflects the permeability of the capillary endothelium and overlying synovial intima, and the surface available for fluid exchanges.! This parameter was used to calculate protein permeability rather than the osmotic reflection coefficient because the latter must be measured at flow independent states.25 To obtain flow independent states, marked increases in venous pressures are required, which would have produced confounding effects such as tissue edema. Because synovial fluid production, and thus lAP were increased in inflamed joints, we measured PS from 300 to 330 minutes, when lAP was at atmospheric pressure, as the value of PS is increased with increased IAP.6,26 Total protein concentration was measured in circuit arterial blood as an estimate of capillary total protein concentration, because we were interested in blood-to-joint protein flux. In addition, for large slowly exchanging molecules such as protein, arterial or mixed venous protein concentrations provide good estimates of capillary concentration.! Our estimates of PS in control joints compare to those

117 previously calculated for the blood-joint barrier of the normal human knee.i Inflamed joints had protein permeability that were raised approximately 5 times that of control joints. A similar observation has been reported for human rheumatoid knees,27 which also reported a greater increase in protein permeability with increased solute dimension.

Synovial membrane permeability to albumin was significantly increased in inflamed joints, and this increase was more significant in the synovial villi region of the joint.

Synovial villi are more vascular and receive a more abundant blood flow than the fibrous synovial membrane.4 Increased albumin clearance has been previously reported in acute articular inflammation, but the effect of synovial membrane type or of innervation had not been examined. The present study illustrates the importance of synovial villi for synovial fluid turnover. Therapeutic synovectomy occasionally performed for the treatment of joint disease may alter clearance of metabolites and synovial fluid kinetics, and these potentially detrimental effects should be considered.

Experimental models of limb isolation demonstrate no significant difference in vascular reactivity between freshly amputated limbs and limbs stored for 12 hours,28 indicating that short term ischemia, such as occurred during surgical preparation of the model, would not contribute to alter vascular responsiveness. In denervated preparations, blood flow would no longer be under control of the sympathetic nervous system, but rather locally by vascular endothelium or metabolic substances. However the lack of effect of denervation to affect the progressive decrease in vascular resistance supports the preservation of vascular reactivity, and the relative importance of local vasomotor function rather than sympathetic innervation to control vascular responsiveness. These findings are in contrast with skeletal vascular responsiveness, where isolated preparations showed a decreased resistance in the immediate post-preparatory period, followed by a progressive increase over tim e.29 However, recordings without treatment were not performed after 75

118 minutes of equilibration, so it is unknown whether vascular resistance would then decrease again with time. These findings may also reflect a different response of skeletal vascular endothelium in comparison with digit vascular endothelium.

Even though attempts were made to minimize trauma during surgical manipulations, there was significant vascular reactivity which appeared to last approximately 2 hrs. Interestingly, but not unexpected, total vascular resistance of the isolated joint was higher in innervated preparation again illustrating the importance of innervation in vascular responsiveness

In conclusion,we characterized the hemodynamic, metabolic, and fluid exchange responses of innervated and denervated joints and their response to inflanunation. The altered permeability observed with denervation illustrates the importance of regional innervation in transsynovial exchanges, as was further corroborated by our findings of significant tissue edema. The significant increase in oxygen demand with inflammation confirms the importance of appropriate oxygen delivery to articular tissue during inflammation, and supports a role of ischemia in the pathogenesis of joint disease.

Beneficial therapeutic intervention that can be gained in arthritis firom such conclusions include improvement of oxygen delivery to the joint by promoting blood flow and minimizing edema formation, and decreasing demand by encouraging joint rest. The increased permeability to small molecules observed with inflammation has direct implications for local therapeutic intervention, and raises the question of extrapolation of pharmacokinetic studies performed in normal joints.

119 Footnotes

a Cardiomax II, Columbus Instruments, Columbus, OH. b Honeywell VRI2, Electronics for Medicine Inc, Pleasantville, NY c Masterflex variable speed pump. Cole Palmer Int, Chicago, EL d Model 7D Polygraph, Grass instruments, Quincy, MA e Cellular Products Inc., Buffalo, NY f Micro-BCA protein assay. Pierce Chemical Co, Rockford, IL g ABL 500, Radiometer, Copenhagen, Denmark h Coherent, Palo Alto, CA i ACAS 570 Interactive laser cytometer. Meridian Instruments, Inc. Okemos, MI j Cytofluor 2350 Fluorescence Measurement System; Astrascan Ltd., Isle of Man, British Isles.

120 Pump Arterial pressure To VR12

FT03 weight a To physiograph Arterial circuit

Reference sample

Venous pressure

Median artery To physiograph Median vein Synovial fluid

Venous collection Venous circuit

Figure 5.1. Illustration of the isolated pump-perfused auto-oxygenated isolated joint model used in the study.

121 Figure 5.2. Circuit arterial pressure (A) and total vascular resistance (B) among groups and across time in isolated metacarpophalangeal joints from Baseline to 60 minutes.

(Group 1 - ■ Control; Group 2-# Control-Denervated; Group 3- A Inflamed; Group 4-

♦ Inflamed-denervated). There was no significant difference among groups.

122 Total vascular resistance Circuit arterial pressure (mmHg) h-ft Is) î I I I I 2 Ê § ë Ê S S § ë %

n 4 >-*+

in u i

è ~

in in

o g - Figure 5.3 Total vascular resistance (TVR) of among groups and across time in isolated metacarpophalangeal joints. (Group I - ■ Control; Group 2-# Control-Denervated;

Group 3- ▲ Inflamed; Group 4- ♦ Inflamed-denervated) There was a significant decrease in TVR over time in all groups (P=0.009), and there was no significant difference detected among groups.

124 Total vascular resistance (mmHg/ml/min) o o o o o o o o o e o o o o e o o o P

1 ^ 11111 1 I 'I' lit' If I ' ' ' I ' ' ' ' I ' ' ' ' I ' ' ' ' I ' ’ ' ' I % Figure 5.4. Synovial membrane blood flow determined by colored microspheres among groups and across time in isolated metacarpophalangeal joints. (Group 1 - ■ Control;

Group 2-# Control-Denervated; Group 3- ▲ Inflamed; Group 4- ♦ Inflamed-denervated)

♦Synovial blood flow was significantly increased (P=0.013) at 60 and 330 minutes compared to baseline and there was no significant difference detected among groups.

126 1400- f 1200 -

1000 -

T3 800-

« 600-

400

^ 200 -

0 50 100 150 200 250300 350 Time (minutes)

Figure 5.4

127 Figure 5^. Synovial fluid production among groups from 240 to 330 minutes in isolated metacarpophalangeal joints.* Indicates a significant difference between inflamed

(groups 3 and 4) and control joints (Groups 1 and 2) (P=O.OOI).

128 Group 1 Group 2 Group 3 Group 4

Control Innervated Control Denervated Inflamed Innervated Inflamed Denervated

Figure 5.5

129 Figure 5.6. Oxygen delivery among groups and across time in isolated metacarpophalangeal joints. (Group 1 - ■ Control; Group 2-# Control-Denervated;

Group 3- ▲ Inflamed; Group 4- ♦ Inflamed-denervated) DO 2 signiflcantly increased across time (P<0.0001). There was no signiflcant difference detected among groups.

130 1.2-1

1 0.8-

0.6 -.

■s 0.4

0.2

0 100 150 200 250 300 35050 Time (minutes)

Figure 5.6

131 Figure 5.7. Oxygen consumption (VO 2) and extraction ratio (0% ER) among groups and across time in isolated metacarpophalangeal joints. (Group I - ■ Control; Group 2-#

Control-Denervated; Group 3- ▲ Inflamed; Group 4- ♦ Inflamed-denervated) * Indicates a significant difference between control (Groups 1 and 2) and inflamed joints (Groups 3 and 4) for VO 2) (P=0.01) and O2 ER (P=0.04).

132 (ml/min/g) O ER (ml/min/g) 2 ooooooooo î booboobbbP (/I Li

4 *

0 0 -

w Figure 5.8. Photomicrographs (20 x) of laser argon acquired images of evans-blue albumin fluorescence of synovial membrane among groups and at fibrous or villous sites, demonstrating depth of absorption. Mean depth of absorption in the villous region for inflamed groups (groups 2 and 4) was 964.85 ± 141.55 |im.

134 CONTROLINFLAMED

FIBROUS

VILLOUS Figure 5.9. FTTC-dextran concentration in circuit plasma effluent among groups and across time in isolated metacarpophalangeal joints from 330 to 360 minutes. (Group 1 - ■

Control; Group 2-# Control-Denervated; Group 3- ▲ Inflamed; Group 4- ♦

Inflamed-denervated) * Indicates a significant difference between control (Groups I and 2) and inflamed joints (Groups 3 and 4) (P=0.02).

136 .2 1.4

€ 0.6

M 0.2

1 I I I r~ I I I I 10 12 14 16 18 20 22 25 30 Time (minutes)

Figure 5.9

137 Baseline 60 minutes 120 minutes 180 minutes 240 minutes 300 minutes 330 minutes

Cardiac output (l_/mln/kg) Group 1 (Control-Innervated) 0.103 ±0.008 0.091 ± 0.006 0.066 ± 0.007 0.064 ±0.009 0.062 ± 0.009 0.079 ± 0.011 0.061 ± 0.011 Group 2 (Control-Denervated) 0.110 ± 0.02 0.1 ± 0.02 0.067 ± 0.01 0.067±0.014 0.063±0.016 0.063±0.013 0.064±0.013 Group 3 (Inflamed-lnnervated) 0.100±0.009 0.093 ± 0.005 0.093 ± 0.007 0.066 ± 0.006 0.091 ± 0.006 0.063 ± 0.004 0.076 ± 0.004 Group 4 (Inllamed-Denervated) 0.066 ± 0.016 0.057 ± 0.012 0.057 ± 0.011 0.065 ± 0.015 0.060 ± 0.013 0.057 ± 0.013 0.056 ± 0.014 Overall 0.094 ± 0.007 0.064 ± 0.006 0.060 ± 0.005* 0.060 ± 0.006* 0.078 ± 0.006* 0.074 ± 0.006* 0.074 ± 0.006* Mean arterial pressure (mtnHg) Group 1 (Control-Innervated) 120.7 ± 10.7 123.6 ± 11.5 127 ± 6.1 126.5 ± 7.7 124.3 ± 7.6 120.3 ± 9.5 118.3 ± 7.4 Group 2 (Control-Denervated) 98.7 ± 14.9 117 ± 8.1 116.7 ± 11.2 115.5 ± 12.5 106.7 ± 10.4 111 ± 11.6 113 ± 12.1 Group 3 (Inllamed-lnnen/ated) 114.0 ± 9.3 114.3 ± 7.5 114.5 ± 5.6 102.3 ± 11.3 111.8 ± 6.1 104.7 ± 8.4 100 ± 12.6 Group 4 (Inllamed-Denervated) 111.8 ± 111.4 121.7 ± 6.3 112.7 ± 6.6 103.6 ± 11.3 101.6 ± 11.5 105.1 ± 10.4 97.6 ± 1 1 .6 Overall 111.2 ± 5.7 119.3 ± 4.2 117.5 ± 3.7 111.6 ± 5.5 111.2 ± 4.7 110.1 ± 4.9 106.6 ± 5.5 Systolic arterial pressure (mmHg) Group 1 (Control-Innervated) 153.5 ± 14.2 156.6 ± 14.9 157.7 ± 10.1 161.2 ± 10.7 155.7 ± 10.7 151.5 ± 11.5 153 ± 11.2 Group 2 (Control-Denervated) 137.6 ± 15.6 148.6 ± 10.2 155 ± 13.5 144.6 ± 14.5 136.5 ± 15.9 141.3 ± 15.9 136.3 ± 17.6 Group 3 (Inllamed-lnnervated) 140.2 ±7.6 145.5 ± 9.2 145.8 ± 8.3 109.6 ± 22.5 143 ± 6.6 132.6 ±8.6 125.2 ± 12.4 Group 4 (Inllamed-Denervated) 142.3 ± 12.4 146.6 ± 6.7 123 ±19.7 129.3 ±12.4 126.9 ± 11.4 129.7 ± 11.3 120.6 ± 15.1 Overall 143.4 ± 6.1 149.9 ± 5.2 144.5 ± 7.3 136.0 ± 6.2 141.0 ± 5.6 136.5 ± 5.9 133.7 ± 7.2 Systemic Pa02 (mmHg) Group 1 (Control-Innervated) 311.1 ± 31.1 291.4 ± 37.5 264.9 ± 40.7 276.2 ± 42.3 216.6 ± 36.2 294.2 ± 44.4 305.7 ± 36.6 Group 2 (Control-Denervated) 241.3 ± 13.9 209.9 ± 20.3 199.0 ± 22.7 190.4 ± 25.6 235.0 ± 45.7 153.9 ± 30.3 149.5 ± 32.2 Group 3 (Inllamed-lnnervated) 169.6 ± 36.3 146.9 ± 35.5 146.9 ± 35.7 150.6 ± 33.3 196.9 ± 42.3 160.1 ± 52.5 161.7 ± 35.6 Group 4 (Inllamed-Denervated) 226.9 ± 41.0 191.7 ± 45.1 216.1 ± 50.6 211.1 ± 41.4 196.6 ± 46.9 250.9 ± 27.9 263.6 ± 36.5 Overall 237.3 ± 16.5 209.3 ± 20.2 212.9 ± 21.2 207.4 ± 19.5 207.6 ± 19.7 221.1 ± 21.5 221.9 ± 21.5 Systemic PaC 02 (mmHg) Group 1 (Control-Innervated) 54.6 ± 7.3 56.4 ±7.7 40.9 ± 1.3 37.9 ±2.9 42.3 ±2.6 42.4 ± 4.5 43.6 ± 5.1 Group 2 (Control-Denervated) 37.2 ± 1.0 46.4 ± 2.9 43.1 ±3.7 39.4 ± 0.5 40.0 ± 1.4 44.5 ± 1.4 42.3 ± 3.0 Group 3 (Inllamed-lnnervated) 43.2 ±2.6 39.7 ± 2.9 40.6 ± 1.9 44.6 ± 2.4 40.4 ± 2.1 40.3 ± 2.7 46.6 ± 1.6 Group 4 (Inllamed-Denervated) 47.2 ± 4.7 50.9 ± 5.6 45.6 ±2.0 43.5 ± 2.9 44.6 ± 2.7 47.3 ± 3.2 46.5 ± 4.0 Overall 45.6 ± 3.9 46.4 ± 4.6 42.5 ± 2.2 41.4 ± 2.2 41.9 ± 2.2 43.6 ± 2.9 45.3 ± 3.5

Table 5.1 Systemic hemodynamics and blood gas analysis (mean ± SE) among groups in horses submitted to isolated joint preparation. * indicates a significant difference from baseline TIME (minutes) Baseline 60 minutes 120 minutes ISO minutes 240 minutes 300 minutes 330 minutes Circuit arterial flow (ml/mln) Overall 14.29 15.88 17.17 18.42 23.77* 23.15* 27.24* ± 4.64 ± 5.38 ± 5.05 ± 5.96 ± 6.89 ± 6.65 ± 8.06 Group 1 (Control-Innervated) 13.00 15.42 14.00 14.92 15.67 17.67 21.26 ± 7.86 ± 8.62 ± 8.42 ± 8.86 ± 8.39 ± 8.23 ± 7.73 Group 2 (Control-Denervated) 12.42 13.92 15.17 15.17 22.67 16.50 25.67 ± 2.96 ± 3.87 ± 2.63 ± 4.52 ± 6.99 ± 4.49 ± 8.50 Group 3 (Inllamed-lnnervated) 13.88 13.05 13.38 12.88 22.05 27.72 27.89 ± 2.94 ± 3.69 ± 4.11 ± 5.84 ± 6.79 ± 8.75 ± 9.40 Group 4 (Inllamed-Denervated) 17.86 21.14 26.14 30.71 34.71 30.71 34.14 ± 4.80 ± 5.34 ± 5.04 ± 4.62 ± 5.37 ± 5.13 ± 6.61 Circuit arterial pressure (mmHg) Overall 151.45 148.23 150.74 150.20 152.29 150.95 153.02 ± 2.42 ± 7.03 ± 4.35 ± 3.48 ± 4.88 ± 3.79 ± 5.20 Group 1 (Control-Innervated) 152.50 145.50 151.83 159.67 152.00 145.83 153.67 ± 2.74 ± 7.14 ± 3.31 ± 3.77 ± 4.32 ± 2.75 ± 8.03 Group 2 (Control-Denervated) 150.83 148.67 147.83 149.00 149.33 149.67 149.67 ± 1.08 ± 1.78 ± 6.49 ± 2.73 ± 2.92 ± 1.87 ± 2.97 to Group 3 (Inllamed-lnnervated) 150.33 157.33 154.00 148.00 148.67 151.17 151.17 ± 1.48 ± 9.15 ± 2.39 ± 2.14 ± 2.78 ± 1.89 ± 4.08 Group 4 (Inllamed-Denervated) 152.14 141.43 149.29 144.14 159.14 157.14 157.57 ± 2.19 ± 11.69 ± 4.48 ± 2.81 ± 7.52 ± 5.77 ± 2.85 Circuit venous pressure (mmHg) Overall 10.12 10.87 10.21 10.23 9.85 9.90 11.17 ± 0.40 ± 1.04 ± 0.85 ± 0.52 ± 0.56 ± 0.56 ± 1.01 Group 1 (Control-Innervated) 10.67 10.17 10.00 9.67 10.00 9.83 10.83 ± 0.49 ± 0.79 ± 1.10 ± 0.21 ± 0.68 ± 0.54 ± 1.11 Group 2 (Control-Denervated) 9.83 12.17 10.33 10.17 9.83 10.00 10.67 ± 0.31 ± 1.54 ± 0.21 ± 0.65 ± 0.48 ± 0.45 ± 0.71 Group 3 (Inllamed-lnnervated) 9.86 10.29 10.29 10.86 9.71 9.86 12.00 ± 0.40 ± 0.78 ± 1.25 ± 0.70 ± 0.52 ± 0.67 ± 1.21 Group 4 (Inllamed-Denervated) 10.12 10.87 10.21 10.23 9.85 9.90 11.17 ± 0.40 ± 1.04 ± 0.85 ± 0.52 ± 0.56 ± 0.55 ± 1.01

Table 5.2 Local hemodynamics (mean ± SE) among groups and across time in isolated joints. * indicates a significant difference from baseiine Synovial fluid total oretain fmq/dl) Time Group 1 Group 2 Group 3 Group 4 Overall Mean SE Mean æ Mean £ Mean £ Mean £ Baseline 2.15 0.15 2.08 0.08 2.17 0.17 2 .00 0.00 2.10 0 .10 240 minutes 2.57 0 .2 4 3 .30 0.30 3.37 0.37 3.14 0.47 3.09* 0 .35 300 minutes 3.03 0.42 3.20 0.28 3.27 0.52 3.49 0 .47 3.25* 0 .4 2 330 minutes 3.15 0.38 3.57 0.33 4.25 0.44 3 .97 0.45 3.73* 0 .40

Synovial fluid pH Overall Time Group 1 Group 2 Group 3 Group 4 Mean £ Mean SE Mean S Mean SE Mean £ Baseiine 7.38 0.02 7.35 0.04 7.38 0.00 7.3 4 0.00 7.36 0 .0 2 240 minutes 7.20 0.02 7.20 0.02 7.17 0.13 7.2 7 0.03 7.21* 0 .05 300 minutes 7.37 0 .00 7 .3 2 0.05 7.41 0.02 7.28 0.06 7.35 0.03 330 minutes 7.48 0.00 7.29 0.00 7.14 0.00 7.34 0.10 7.31 0 .0 2

Synovial fluid Pa02 (mmHg) Time Group 1 Group 2 Group 3 Group 4 Overall Mean S Mean SE Mean £ Mean £ Mean £ Baseline 139.95 5.85 87.76 7.56 99.50 0.00 130.60 0.00 114.45 3 .35 240 minutes 46.27 5.63 71.23 4.91 51.90 19.60 9 5 .4 7 15.50 66.21* 11.41 300 minutes 57.50 0.00 107.80 34.44 129.20 41.90 6 2 .4 5 5.85 89.24 20.55 330 minutes 151.20 0.00 73.80 0.00 33.30 0.00 156.97 88.47 103.82 22.12

Synovial fluid PaC02 (mmHg) Time Group 1 Group 2 Group 3 Group 4 Overall Mean æ Mean S Mean £ Mean £ Mean £ Baseiine 48.85 0.63 48.71 2.88 48.60 0.36 47.02 1.22 48.30 1.27 240 minutes 66.17 1.17 71.34 7.08 66.77 3.70 59.31 2.82 65.90* 3.69 300 minutes 52.78 0.60 53.55 3.77 51.97 1.19 54.94 1.42 53.31 1.74 330 minutes 45.45 1.25 40.36 8.12 51.17 4.47 49.39 5.29 46.59 4 .78

Synovial fluid 02 gradient (mmHg ) Time Group 1 Group 2 Group 3 Group 4 Overall Mean SE Mean æ Mean £ Mean £ Mean £ Baseiine 195.95 32.35 151.10 14.29 73.87* 33.42 129.14*¥ 35.41 173.53 2 8 .8 7 240 minutes 226.67 42.72 127.28 34.02 82.25* 41.34 121.08*¥ 28.23 176.98 36.58 300 minutes 205.27 43.02 59.93* 23.14 79.77* 48.32 164.68*¥ 22.49 205.27 34.24 330 minutes 177.00 40.39 52.55* 23.63 71.32* 29.80 124.37*¥ 35.48 177.00 32.32

Permeability surface area product (ml/mln) Group 1 Group 2 Group 3 Group 4 Overall Mean SE Mean Mean £ Mean £ Mean £ 330-330 minutes 0.00057 0.00014 0.00036 0.00036 0.0039 0.0015 0.0037 0.0003 0.0021 0.0005

Intraarticular pressure (mmHg) Group 1 Group 2 Group 3 Group 4 Overall Mean SE Mean SE Mean £ Mean £ Mean £ 240 minutes 1.08 a 0.27 1.33 a 0.80 7.00 b 3.17 6.00 b 1.70 3.85 1.49

Table 5.3 Synovial fluid total protein. pH, Pa02, PaC 02.0 2 gradient, permeability surface area product and intraarticular pressure among groups and across time In isolated Joints * indicates a significant diflercnce from baseline ¥ indicates a significant difference between group 3 and group 4 for that time period different letters are significantly different

140 Wet/dry ratio Group 1 (Control-Innervated) 1.02 ± 0.03 Group 2 (Control-Denervated) 1.11 ± o.ot Group 3 (Inflamed-lnnervated) 1.06 ± 0.02 Group 4 (Inflamed-Denervated) 1.12 ± 0.03t

Table 5.4 Synovial membrane wet/dry ratio among groups in isolated joints. t indicates a significant difference between innervated (Groups I and 3) and denervated (Groups 2 and 4) joints. (P=0.CX)9)

141 Fibrous synovium Viilous synovium Depth of fluorescence Group 1 (Control-innervated) 493.3 ± 83.5 685.7 ± 96.7* Group 2 (Control-Denervated) 615.6 ± 229.5 774.0 ± 143.1* Group 3 (Inflamed-lnnervated) 489.0 ± 113.2 1075.0 ± 112.7*¥ Group 4 (Inflamed-Denervated) 433.7 ±1010 854. 7 ± 170.4*¥

Table 5.5 Depth of Evans blue-albumin fluorescence (p.m) among groups in isolated equine joints *indicates a significant difference between villous and fibrous synovium (P=0.000 ¥ indicates a significant difference between inflamed (groups 3 and 4) and control (groups 1 and 2) joints (P=0.023).

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15. Coderre, T. J., A. I. Basbaum, J. D. Levine. Neural control of vascular permeability: interactions between primary afferents, mast cells and sympathetic efferents. JNeurophys 62:48-58, 1989.

143 16. Miao, F. J.-P., W. Janig, J. D. Levine. Role of sympathetic postaganglionic neurons in synovial plasma extravasation induced by bradykinin. JNeurophys 75:715-724, 1996.

17. Konttinen, Y. T., P. Kemppinen, M. Segerberg, et al. Peripheral and spinal neural mechanisms in arthritis, with particular reference to treatment of inflammation and pain. Arthritis Rheu 37:965-982, 1994.

18. Lembeck, P., P. Holzer. Substance P as neurogenic mediator of antidromic vasodilation and neurogenic plasma extravasation. Arch Pharmacol 310:175-183, 1979.

19. Schumacker, P. T., S. M. Cain. The concept of critical oxygen delivery. Intensive Care Med 13:223-229, 1987.

20. Strand, E., G. S. Martin, M. P. Crawford, et al. Intra-articular pressure, elastance, and range of motion in flexion in the equine metacarpophalangeal joint. Am J Vet Res 56:1362-1371, 1994.

21. Blake, D. R., J. Uns worth, J. M. Outhwaite, et al. Hypoxic-reperfusion injury in the inflamed human joint. The Lancet 289-292, 1989.

22. Woodruff, T., D. R. Blake, J. Freeman, et al. Is chronic synovitis an example of reperfusion injury? Ann Rheum Dis 45:608-611, 1986.

23. Bertone, A. L., J. Hardy, E. J. Simmons, et al. Physiologic responses of a novel isolated joint model. Abstr, Orthop Res Soc 1994.

24. Hardy, J., A. L. Bertone, W. W. Muir. Pressure-volume relationships in normal equine midcarpal joints. JAppl Physiol 78:1977-1984, 1995.

25. Taylor, A. E. Capillary fluid filtration. Starling forces and lymph flow. Circ Research 49:557-575, 1981.

26. Levick, J. R. The influence of hydrostatic pressure on trans-synovial fluid movement and on capsular expansion in the rabbit knee. J Physiol (Lond) 289:69-82, 1979.

27. Kushner, I., J. A. Somerville. Permeability of human synovial membrane to plasma protein. Arthritis Rheum 14:560-570, 1971.

28. Ono, H., Y. Nakagawa, S. Mizumoto, et al. Evaluation of vascular compliance and vasoconstrictive reactions in amputated hindlimbs of rat. J Orthop Res 13:375-381, 1995.

29. Mellander, S., M. Maspers, J. Bjomberg, et al. Autoregulation of capillary pressure and filtration in cat skeletal muscle in states of normal and reduced vascular tone. Acta Physiol Scand 129:337-351, 1987.

144 CHAPTER 6

CELL TRAFFICKING, MEDIATOR RELEASE AND ARTICULAR CARTILAGE METABOLISM IN ACUTE ARTHRITIS OF INNERVATED OR DENERVATED ISOLATED JOINTS

Introduction

Acute articular inflammation is clinically recognized by joint effusion, periarticular edema, local heat and pain, lameness of the affected limb, and decreased range of motion of the joint. 1 Routine synovial fluid analysis in acute articular inflammation reveals variable degrees of neutrophilic leukocytosis, increased protein concentration, and decreased mucin precipitate quality.! Synovitis accompanies most forms of joint disease, and predominates in acute articular inflammation.^ Inflamed synovial cells can release mediators such as interleukin-1 and prostaglandin E 2 (PGEi)^ which in turn activate adjacent synoviocytes,^ and alter the balance of chondrocyte matrix synthesis and degradation towards a degradative pathway .5

The investigation of acute articular injury is important because single articular trauma can lead to progressive degenerative joint disease if not identified or treated appropriately. Therapeutic efforts may be more successful to restore joint health in acute injury, if gross changes have not occurred. Several models of acute articular inflammation have been developed in equids, including intra-articular injection of endotoxin,^ carrageenan,? amphotericin B,8 and bacteria.9 These models provide important clinical and clinicopathologic information that allowed the investigation and comparison of therapeutic

145 modalities, however, the acute dynamic events including cellular migration and mediator release occurring in acute inflammation were not documented.

Recent advances in joint disease emphasized an important role of innervation in modulating articular inflanunation. 10-12 Clinical and experimental data suggest that a decreased inflammatory response occurs in denervated joints. Antidromic release of neuropeptides, particularly substance P, from small C fibers, has been shown to occur in articular inflammation,!3 and neuropeptide depletion has been associated with improvement in clinical signs of disease. 14 Increased substance P concentrations have been measured in synovial fluid of horses with joint disorders, 15 and substance P staining nerve fibers have been identified in equine synovial membrane. 15-17 In vitro studies have indicated a synergistic effect of substance P and interleukin-1 to degrade articular cartilage.I 8 Despite this information, no experimental or clinical information is available on the effect of denervation to modulate articular inflammation in the horse. Investigations of the early events in articular inflammation, and the possible role of neural modulation needed to be performed in an appropriate model.

An isolated joint preparation model for the study of joint physiology, 19 has been used to characterize the permeability, fluid exchange and oxygen metabolism in acutely inflamed, innervated or denervated isolated joints .20 We documented increased permeability, oxygen metabolism and oxygen debt with acute inflammation, and an effect of denervation to decrease articular permeability to large molecular weight molecules and increase edema formation.

The purpose of the present study is to describe the cellular events (synovial and chondrocytic), cytokine, eicosanoid and neuropeptide release in acute inflammation of innervated or denervated isolated joints, and associate these events with the physiologic alterations of innervated or denervated joints.

146 Materials and methods

Horses -Twenty-four adult horses (body weight 330-550 kg, 8 females, 8 geldings, 8 stallions) of mixed breeds were used. Horses were determined free of metacarpophalangeal joint disease by thorough clinical examination, synovial fluid analysis, and gross post-mortem examination. All experiments were conducted in accordance with the Animal

Care and Use Guidelines of The Ohio State University.

Experimental procedures

Instrumentation

Horses were instrumented for hemodynamic measurements» and intravenous drug administration prior to induction of general anesthesia as previously described (see Chapter

5).

Anesthesia and monitoring

Only one horse was studied on a given day. Horses were sedated and anesthesia was induced as previously described (see chapter 5). Anesthesia was maintained with sodium pentobarbital (loading dose 4 mg/kg, maintenance 5-15 mg/kg/hour, iv, to effect) and controlled ventilation with 100% oxygen. A bipolar ECO (base-apex lead) was recorded throughout each study for determination of heart rate and rhythm. Arterial pressure and ECG were continuously displayed and semicontinuously recorded on a multichannel recorder^. Systemic arterial blood gases, packed cell volume and total protein were determined hourly. Intravenous polyionic isotonic fluids were administered (5-10 ml/kg/hr) throughout each study. Mean systemic arterial pressure was maintained at > 70

147 mm Hg, and ventilation was adjusted to maintain PCO 2 < 55 mmHg. Heparin (50,000 lU iv) was administered every 90 minutes.

Joint isolation procedure and instrumentation (Fig. 6.1)

One metacarpophalangeal joint was randomly selected and the horse was positioned with the selected limb lowermost. All circuit tubing for circulation through precalibrated pumpsc was thoroughly flushed with heparinized saline. The MCP was isolated as previously described (see Chapter 5). In innervated preparations, the medial and lateral palmar nerves were isolated and preserved. The isolated joint was suspended from a calibrated FT03 transducer connected to a physiography for continuous measurement of weight changes. All circuit pressure transducers were placed at the level of the joint preparation. Initial circuit arterial flow was set at 10 ml/min until completion of the preparation and establishment of isogravimetric state (see below). This joint isolation technique established a pressure-driven auto-oxygenated, pump-perfused, extracorporeal isolated joint model.

Experimental protocol

The joint preparation model was randomly assigned to one of four groups:

1-Control; 2) Control-denervated; 3)Inflamed; 4)Inflamed-denervated. In groups 3 and 4, inflammation was induced by intraarticular injection of 0.35 ng/kg BW of recombinant human interleukin-1 Be (D-1B) .After complete joint isolation and instmmentation, isogravimetric state (no gain or loss of weight in the isolated preparation) was established and maintained for 20 minutes by adjustment of arterial flow. In all groups, the preparation was maintained for 360 minutes, after which horses were euthanized with an overdose of pentobarbital (1(X) mg/kg, iv), and specimens collected. Baseline (time 0) was considered

148 as the time after establishment of isogravimetric state, collection of the baseline synovial fluid sample, and before injection of 1116 in Groups 3 and 4.

Blood sample collection

Circuit arterial and venous blood were collected at baseline and hourly thereafter, in heparinized syringes, and transported on ice for blood gas determination, and measurement of packed cell volume and total protein (refractometer).

Synovial fluid collection and analysis

Synovial fluid was collected at time 0, and at 240,300 and 330 minutes. Synovial fluid volume was recorded from 240 to 300 minutes and from 300 to 330 minutes by collection into a graduated cylinder. Fluid for clinical pathological analysis was collected into 2 ml evacuated tubes containing 1.5 mg of K 3 EDTA/mlf The total number of nucleated cells was determined by use of an automated hematology analyzerg. Differential leukocyte counts were made by counting 1 0 0 cells on direct smears that were air dried and stained with a Wright-Giemsa stainh- Synovial fluid samples for the analysis of the cytokines H-Ip, and interleukin 6 (11-6) were collected in Eppendorf tubes and centrifuged, and the cell-free supernatant was stored at -20°C for 24 hrs then at -80°C until assay.

Cytokine assays were performed using commercially available ELISA kits'. These assays employ the quantitative ‘sandwich’ enzyme immunoassay technique, using monoclonal antibodies raised against human cytokines, as well as human recombinant cytokines as standards. Sensitivity of the assays were 1.3 pg/ml and 3.2 pg/ml for II-ip, and H- 6 , respectively. Synovial fluid for the PGE 2 assay was collected in chilled Eppendorf tubes containing 25 pi of 100 mM Na 2EDTA and 25 pi of 10 mM meclofenamic acid. Tubes were centrifuged for 1 0 minutes, and the cell-free supernatant was collected and stored at

149 -20°C for 24 hours, then at -80°C until assayed. PGE 2 concentration was determined by use of commercially available PGE 2 EIA kits). Absorbance was read at 405 nm by use of a microplate reader. Absorbance was correlated with concentration by use of a standard curve ranging from 1 0 to 5000 pg/ml. Samples with >5000 pg/ml PGE 2 concentrations were reanalyzed after dilution. Synovial fluid for substance P determination was collected in chilled Eppendorf tubes and centrifuged at -4°C. The cell-free supernatant was collected, placed in Eppendorf tubes, immediately frozen in liquid nitrogen and stored at -80°C until assay. Substance P concentrations were quantified in duplicate, using standard radioimmunoassay techniques. The primary antibody was used as a dilution of 1:10,000.

Standard curves were constructed using i25i-radioIabelled ligand/lOOpl (5,000 counts per min) and increasing concentrations of unlabeled standard ligands for the SP assays.

Concentration ratios for IIIB and 116 were calculated using group 1 as the reference group to assess relative differences. PGE 2 results for each time within each group are expressed as a ratio to baseline values because marked variations in baseline values existed, as previously reported.? Interleukin-16, interleukin - 6 and substance P results are expressed as concentrations (pg/ml), content (pg) and ratio. Content was calculated by multiplying the average of concentrations at 240 and 300 minutes and 3(X) and 330 minutes by volume of synovial fluid collected between 240 and 300 minutes, and 300 and 330 minutes. Total content for these 90 minutes was obtained by adding these values.

Synovial membrane and cartilage collection

The dorsal aspect of the isolated MCP joint and the contralateral MCP joint were aseptically prepared. FTTC-dextran and Evans blue albumin were injected into the isolated and contralateral joint 30 minutes before euthanasia. After euthanasia, the preparation was disassembled and weighed, to expose articular cartilage to the same treatments as that in the

150 isolated joint (see chapter 5). Using aseptic technique, the dorsal skin and joint capsule were incised and the joint cavity was exposed. The joint was carefully inspected for evidence of cartilage erosions, synovial membrane proliferation or presence of osteophyte.

The presence of gross abnormalities excluded the preparation from the study.

Svnovial membrane collection: Three synovial membrane and underlying joint capsule specimens ( 2 0 mm x 1 0 mm each) were collected from both joints and placed on a wooden support to maintain shape. One specimen was placed in fixative (4ml 25% glutaraldehyde, 25 ml 0.2M sodium phosphate buffer, 21 ml water, 1.7 g sucrose) for 24 hours, followed by storage in 70% alcohol, for histopathology and substance P immunostaining. These specimens were processed and stained within 2 weeks of collection. One specimen was packed in OCT, frozen in liquid nitrogen and stored at

-70°C. This specimen was processed for fluorescence studies (see chapter 5). One specimen was used for determination of wet/dry ratio and microsphere determination (see chapter 5).

Cartilage collection Full-thickness cartilage specimens were obtained from the medial aspect of the sagittal ridge of each distal metarcarpi using a 2 mm diameter bone biopsy trocar. Care was taken to collect cartilage from the non-weight bearing aspect of the joint surface, and to limit collection to within 0.5 cm of the cartilage edges. Cartilage specimens from each joint were placed in sterile DMEM medium and transported on ice to the laboratory. Under a laminar flow hood, approximately 100 mg of pooled cartilage from each joint were aliquoted to each of 4 wells of a six well plate: Well 1-18 hour 35[S] incorporation; control joint; Well 2-18 hour 35[S] incorporation; isolated joint; Well 3-

Cartilage degradation study; control joint; Well 4- Cartilage degradation study; isolated joint.

151 Cartilage proteoglycan synthesis and degradation

Cartilage synthesis

One well from each joint was used for synthesis studies. In each well, 2.5 ml of standard medium (DMEM with 10% fetal calf semm, 100 |ig/ml penicillin and 100 pg/ml gentamicin) and 40 pCi of 35[S] were added. Cartilage specimens were incubated for 18 hours at 37 °C in 100% humidity and 5% CO2. After 18 hours, cartilage specimens were placed in a separate vial and extracted for 48 hours at 4°C with 4 ml guanidine-HCl in 0.1

M Na-acetate (pH 5.8) containing protease inhibitors (1(X) mM 6 -aminohexanoic acid, 10 mM Na2EDTA, 5 mM benzamidine). Extracted samples were centrifuged for 5 minutes at

4°C to remove unextractable material and dialyzed against deionized water at 4°C for 24 hours (membrane cut-off: 6,000-8,(XX)) to remove unincorporated radiosulfate and to allow for reaggregation of the proteoglycan with endogenous hyaluronan. An aliquot (1(K) pi) of each extract was counted on a scintillation counter, and samples were stored at -70°C until assay for proteoglycan and glycosaminoglycan concentrations. Total protein in each sample was measured by the bicinchoninic acid assay kiti, with bovine serum albumin as standard.

Proteoglycan synthesis was expressed as CPM/pg protein. Endogenous proteoglycan concentration was measured by determination of uronic acid concentration determined on cartilage extracts by the carbazole-borosulfuric acid method using glucuronolactone as standard.21 Proteoglycan concentration is expressed in pg/pg protein. Total glycosaminoglycan concentration was measured by dimethylmethylene blue assay2203 followed by sequential digestion with chondroitinase ABC and keratanasek, to measure chondroitin sulfate and keratan sulfate content, respectively. Chondroitin sulfate was used to construct standard curves. Glycosaminoglycan concentrations are expressed in pg/pg protein.

152 Cartilage degradation

One well from each joint was used for degradation studies. In each well, 2.5 ml of standard medium (DMEM with 10% fetal calf serum, 100 p.g/ml penicillin and 100 |ig/ml gentamicin) and 40 p.Ci of 35[S] were added. Cartilage specimens were incubated for 72 hours at 37 °C in 100% humidity and 5% C02. After 72 hours, half of each cartilage specimen was collected, placed in a separate vial, and extracted as described above. The remainder of the cartilage specimens were rinsed three times in PBS and further incubated in standard medium without radioactive material. Medium was collected and exchanged every 24 hours for 72 hours. After 72 hours (144 hours total incubation), cartilage was placed in a separate vial and extracted as described above. All cartilage and culture medium were dialyzed against water to remove unbound sulfur. Ail dialysis volumes were recorded. Scintigraphic counts were obtained on aliquots of cartilage extracts obtained at 72 and 144 hours, and on daily collected culture medium. Proteoglycan concentrations in daily collected media were determined using the carbazole-borosulfuric acid method (see above). Newly synthesized proteoglycan release in media are expressed as CPM for each time, and as a ratio of experimental to control joints. Endogenous proteoglycan released in media are expressed as jig, and as a ratio of experimental to control joints. The percentage of total proteoglycan (jig) and newly labelled proteoglycan (CPM) released from the cartilage explants into culture media was expressed as foUows24:

% release at time X = jig or CPM of PG in medium at time X xlOO

sum (jig or CPM) of PG in all collected media + jig or CPM of PG in cartilage extract at 144 hrs

Each value was expressed as a percentage and as a ratio to the contralateral limb.

Substance P and Nerve Specific Enolase (NSE ) immunostaining

153 Synovial membrane specimens were fixed, paraffin embedded sectioned and processed for immunostaining as follows. After deparrafinization in xylene, specimens were rehydrated in decreasing concentrations of ethanol, followed by 3% peroxide to block endogenous peroxidase. Specimens were then coated with normal goat semm to block non-specific binding, followed by coating with the primary antibody (rabbit anti-substance

Pm or rabbit anti-NSE) and incubated for 1 hour in a humidified chamber to prevent specimen drying. After rinsing with PBS, the secondary antibody (anti-rabbit biotynilatedn

)was added and incubated for 20 minutes. After rinsing with PBS the tertiary antibody

(streptavidin-peroxidase complex) was added to bind to the secondary antibody and incubated for 20 minutes. The specimens were then coated with freshly prepared diamonobenzidine-peroxide substrate for reaction with the tertiary antibody to produce the brown color. Specimens were dehydrated in increasing concentrations of ethanol, and coverslip mounted.

For each specimen, a blank was prepared where the rabbit anti-neuropeptide was omitted.

Slides were examined under oil immersion and 10 adjacent high powered fields were scrutinized in the fibrous and villous synovium specimens. The number of positive staining nerve filament per field was recorded. The location (perivascular or interstitium) was also recorded.

Histopathology scores

Synovial membrane specimens collected from the dorsal aspect of the experimental and contralateral control metacarpophalangeal joints were fixed, paraffin embedded, sectioned and stained with hematoxylin and eosin for histopathologic examination.

Specimens were evaluated at 20X and lOOX (oil immersion) by three investigators blinded as to the specimen examined. Semi-quantitative morphology scores were assigned on villous and fibrous sections of synovial membrane for extent of edema, hemorrhage,

154 neutrophilic vasculitis, distribution of infiltrate and morphology of the intima in both the villous and the fibrous regions of the synovial membrane. Scores were assigned for each category (see table 6.1). A interobserver mean score was obtained for each category and each site.

Statistical analysis

A three factor analysis of variance (model (innervated or denervated), treatment (control or inflamed) and time) with repeated measures (time) was used to test for differences among groups and across time of synovial fluid white blood cell count, white blood cell differential, 1116,11-6 and substance P concentrations, content and ratios, cartilage extracts and culture medium scintillation counts, hexuronic acid concentration, cartilage extract glycosaminoglycan values and cartilage degradation ratios. Cartilage extract values were obtained for incubations at 18,72 and 144 hours. Degradation values were obtained at 24,

48 and 72 hours. For all cartilage and culture medium analysis, age of the animal was entered in the analysis as a covariate. Histopathology scores were compared with a three way analysis of variance comparing the factors model, treatment and joint (experimental or contralateral control). Synovial fluid PGEi concentrations were expressed as a ratio of baseline samples. Natural logarithmic and arcsine transformations25 were used to stabilize variances of data for white blood cell total count and differential, respectively, before analysis. For all analysis, if a significant F test was obtained, a post-hoc Neuman-Keuls multiple comparison test was used to detect difference in means among groups. A significance level of P<0.05 was assigned for all tests. A coefficient of repeatability among observers was calculated for histopathology scores by doubling the standard deviation of the differences between scores and mean.26

155 Results

Systemic hemodynamics

Systemic mean arterial pressure and PaCOi were maintained within stated limits of mean arterial pressure >70 mmHg and PaC02 < 55 mmHg (See chapter 5). There were no difference detected in systemic hemodynamics or blood gas analysis measurements among groups.

Synovial fluid analysis

All baseline synovial fluid WBC and nucleated cell differential counts were within accepted normal ranged? (total nucleated count < 1.0 x 103 cells/p.1, neutrophils < 10%).

Synovial fluid WBC in groups 1 and 2 significantly increased across time over baseline, with a peak mean WBC of 2.3 x 103 cells/p.1 at 240 minutes. IllB-treated isolated joints

(groups 3 and 4) had significantly greater synovial fluid WBC than control isolated joints

(groups 1 and 2) (Fig 6.2) (P=0.001). Mean synovial fluid WBC in joints of groups 3 and

4 peaked at 330 minutes and was 9.5 x 103 + 2.2 cells/p.1. Maximal WBC in inflamed joints was 26.9 x 103 cells/pl, and was observed at 330 minutes. Synovial fluid WBC differential was within normal reported ranges^? at baseline in all isolated joints (Fig 6.3).

In all groups, % neutrophils significantly increased (P<0.001) and % large and % small mononuclear cell significantly decreased over time. There was no significant effect of treatment or denervation on synovial fluid nucleated cell differential.

Synovial fluid interleukin-1, interleukin-6, PGE 2 and substance P concentrations

Interleukin IB concentration was significantly greater (P < 0.0(X)1) in groups 3 and

4 compared to groups 1 and 2 up to 300 minutes after intraarticular injection of

156 interleukin-16 (Table 6.2). Similarly, DIB content was greater in groups 3 and 4 than groups 1 and 2 (P<0.05) (Table 6.2). When inflamed groups only were considered, DIB concentrations, content and ratio were significantly greater (P=0.04) in group 3 than group

4 at 240,300 and 330 minutes (Fig 6.4 and table 6.2). At 240 minutes, II IB concentrations in inflamed-innervated joints (Group 3) were 19 times greater than control, vs 8.6 times in inflamed-denervated joints (Group 4). In inflamed-denervated joints DIB concentration, content and ratios had returned to levels to significantly different from control by 300 minutes.

Interleukin-6 concentration was significantly greater (P=0.01) in groups 3 and 4 compared to groups 1 and 2 at 240, 300, and 330 minutes (Table 6.3). In contrast to ElB,

D6 concentrations showed an increase across time. Similarly, 116 content was greater in groups 3 and 4 than groups 1 and 2 (P<0.05) (Table 6.3). The concentration ratios of 116 were significantly increased (P=0.01) in inflamed (Groups 3 and 4) compared to control

(Groups 1 and 2) with a peak ratio of 21 times that of the reference group observed at 330 minutes in inflamed-denervated joints (group 4). There was no effect of denervation detected on 116 concentration, content, or ratio.

Synovial fluid PGE 2 ratio significantly increased over time in all groups (P=0.029).

Inflammation (groups 3 and 4) significantly increased PGE 2 ratio over control joints

(groups I and 2) (P=0.038) (Fig 6.6). Synovial fluid from Groups I and 2 had a maximal elevation in PGE 2 of 2.5 ± 0.6 times baseline values whereas inflamed denervated joints

(groups 4) had a mean elevation of 16.9 ± 11.9 times baseline values. These peak ratios were observed at 330 minutes.

Total substance P content in synovial fluid was significantly greater (P=0.025) in groups 3 and 4 than groups 1 and 2 at 300 and 330 minutes (Fig 6.7). There was no

157 significant difference in denervated joints in total substance P amount released in synovial fluid.

Chondrocyte proteoglycan synthesis and content

Proteoglycan synthesis (CPM/p.g protein) and endogenous proteoglycan content

(p.g/|ig protein) and their respective ratios of experimental to control leg were not significantly different among groups or across time (18 hours, 72 hours or 144 hours)

(Table 6.4). Total glycosaminoglycan, chondroitin and keratan sulfate concentrations were not significantly different among groups (Table 6.5).

Proteoglycan degradation

The ratio of newly synthesized proteoglycan released in culture medium was significantly decreased at 24 and 48 hours in Inflamed -denervated group (Group 4)

(P=0.03) (Table 6.6). There was no difference in total uronic acid concentration

(endogenous proteoglycan) released in culture medium among groups or across time (Table

6.7).

Histopathology scores

There was no significant effect of joint isolation, inflammation or denervation on edema scores. Isolated joints (Groups 1-4) had significantly greater (P=0.02) hemorrhage scores than their contralateral control joints. However there was no effect of inflammation or denervation on this score. There was a significant effect of site within the joint, joint isolation and inflammation on neutrophilic vasculitis scores. Villous synovial membrane had greater neutrophilic vasculitis scores than fibrous synovial membrane. Isolated joints had significantly greater neutrophilic vasculitis than their contralateral control joints.

Inflamed joints (groups 3 and 4) had significantly greater (P=0.01) neutrophilic vasculitis

158 scores than control isolated joints (groups I and 2) (Figure 6.8 and table 6.8). There was no effect of denervation on this score. The coefficient of variability among observers was

0.68, indicating that 95% of observations were within a score value of 0.68.

Substance P and NSE immunostaining

Occasional substance P positive peroxidase stained nerve fibers were observed in the interstitium and perivascular space of synovial membrane specimens (Plate 6.2).

Similarly, NSE-positive nerve fibers were observed at those sites. Subjectively, there was a greater density of nerve fibers in the villous area of the synovial membrane compared to the fibrous zone. The number of positive staining fibers were too few and variable to justify quantitative measurements.

Discussion

Interleukin-16 injected in equine joints induced a rapid cellular inflammatory response identified by a synovial fluid leucocytosis at 240 minutes after induction, a significant neutrophilic vasculitis particularly of the villous synovium at 360 minutes after induction, and a significant release of the mediators 116 and PGE 2. Acute denervation during the induction of acute intraarticular inflammation by 1116 resulted in lower increases in 1116, and a more rapid return to baseline values than that observed with innervation. In denervated joints, the ratio of PGE 2 was greater with inflammation than with innervation.

Denervated joints also had less cartilage degradative response to inflammation than denervated joints.

Synovial fluid leukocytosis was observed in HI 6 injected joints. The magnitude of increase in synovial fluid WBC and the neutrophilic response were comparable to that observed in clinical cases of synovitis.^? This model of 1116 (0.35 ng/kg) produced a

159 milder synovitis than that reported after carrageenan or amphotericin B injection where

WBC values are in the 100-150 x 103 cells/p.1 ra n g e J-8 Synovial fluid WBC values after intraarticular endotoxin were dose-dependent.6

Interleukin IB concentrations were significantly higher at 240 minutes (P<0.0005) in inflamed joints, but had returned to baseline values at the end of the study. Mean 11-16 concentrations at 240 minutes in inflamed joints was 19.9 ± 6.5 pg/ml, with a peak value of 85.25 pg/ml. These values agree with the reported range of ElB concentrations obtained in synovial fluid collected from horse with various joint disorders.28 Total ElB content

(Group 3:164.6 ± 46.7 pg/ml. Group 4: 107.19 ± 28.18 pg/ml) were much smaller than the amount originally infused, suggesting that the majority of ElB is cleared by transsynovial pathways or degraded. Control joints had ElB concentrations that were less than 3.0 pg/ml. This single ElB injection presumably did not result in increased or sustained endogenous ElB synthesis, as concentrations continuously decreased following the initial sampling time. The significantly greater ElB in innervated joints suggests a role of sympathetic innervation to contribute to mediator release and inflammation in the joint c a p s u le .2 9

Interleukin 6 concentrations significantly increased (P=0.011) throughout the study period in inflamed joints. Peak E6 values obtained in this study compare well with values obtained in synovial fluid from horses with naturally occurring acute severe joint disorders.28 This increased E6 release may have stemmed from resident synovial cells, chondrocytes, or from the influx of neutrophils.30.3i Previous in vitro studies have reported interleukin 6 release from synoviocytes in response to inflammatory stimuli. In addition, ElB is a direct stimulus for E6 release.3i

The increased PGE; concentrations observed in E IB-inflamed joints is consistent with increased PGE 2 concentrations obtained in clinical joint disease in p e o p le,3 2 and

160 h orses.33 Our values obtained with ElB were lower than that reported with experimentaUy induced carrageenan synovitis in horses, however our inflammation was less.?

Prostaglandin release can be induced by cellular damage, vascular distention or stress, and

E-1. Individual tissues produce a single eicosanoid, and PGE 2 is commonly released from articular inflammation. Equine synoviocytes and chondrocytes have been shown to secrete

PGE2 rather than PGE%. PGE2 has a direct effect on synoviocytes to release mediators.

Variability in control joint PGE 2 concentrations at baseline was probably the result of surgical manipulation because of the variability in baseline concentrations, expression of

PGE2 as a ratio of baseline values aUowed to stabilize variances and to gain a better understanding of patterns of release of PGE 2.

The amounts of substance P released was significantly greater in inflamed joints.

However, there was no significant difference between innervated or denervated joints on substance P release, as might have been expected. The acute denervation in our model may not have influenced the concentration of substance P at the nerve ending, because substance

P is synthesized by the cell body in the dorsal root ganglion, and is carried by axonal transport to the nerve ending where it is stored until release.

Interleukin-16 induced a significant neutrophilic vasculitis in inflamed joints. This response emphasizes the rapidity with which the joint reacts to inflammatory stimulation.

Clinically, pain, effusion and a leukocytic response are observed within hours after exposure to a inflammatory stimuli such as bacteria or trauma. Activation of cell surface adhesion molecules may regulate leukocyte migration and chemotaxis to the joint. Resident articular endothelial and synovial cells can upregulate the expression of adhesion molecule in response to pro-inflammatory cytokines such as interleukin 1.34 At least three distinct adhesion molecules have been described in articular tissue: E-selectin, intercellular adhesion molecule-1 (ICAM-1) and vascular cell adhesion molecule (VCAM-1). E-selectin is

161 probably involved in the initial adhesion of circulating resting neutrophils, and E-selectin was preferentially expressed on small superficial venules in synovium. Our findings further identify a significant neutrophilic vasculitis in the most superficial vessels of the synovium and document the rapidity of the response.34

Our results indicated a degradative response of articular cartilage in irmervated joints exposed to ElB, whereas no effect on synthesis was observed. A significant decrease in degradation of newly synthesized proteoglycan was observed in inflamed denervated joints. In vitro incubation of equine cartilage explants to 111 showed increased degradation and decreased proteoglycan synthesis,24 or decreased synthesis only without effect on degradation.35 However, these latter studies used different labelling times, or did not account for amount incorporated. Measurement of mRNA transcription of degradative enzyme may be more accurate to detect an effect of III. In one report, the degradative response of articular cartilage following exposure to II IB preceded the synthetic effects of the cytokine. Our findings of a short term effect of n IB on cartilage degradation in vitro is consistent with rapid recovery of cartilage following removal of 111.35 The protective effect of denervation on degradation of cartilage supports previous findings of a synergistic response of innervation to enhance the inflammatory response, through neuropeptide release or contributions of postganglionic nerve terminals in the joint capsule to promote mediator release. 11.29

In this model, interleukin-1 was chosen to induce articular inflammation, as it is a reported pro-inflammatory cytokine which has been used in several models of articular inflammation. Interleukin-1 activity is increased in clinical cases of inflammatory arthritis of horses, 36,37 and increased 111 concentrations (ELISA) have been also been reported.28

Human recombinant DIB was used instead of the a isoform, based on previous findings that human recombinant DIB was three to ten times more potent than Ala to stimulate

162 equine synoviocyte PGE 2 production.38 Interleukin-IB has been shown to increase human

PGE2 release by chondrocytes.

This study is the first to report the effects of in vivo intraarticular administration of niB in equine joints. Interleukin-1 has been documented as an important mediator in articular inflammation, and has been isolated from synovial fluid of horses with various forms or articular disease. Incubation of equine cartilage with II-1 in vitro results in a significant inhibition of matrix synthesis and increases matrix degradation. Our study confirms that II-1 can induce a significant inflammatory response in vivo, akin to naturally occurring articular inflammation. Intra-articular injection of recombinant human D la or II IB in rabbit stifle joints resulted in a significant dose-dependent increase in intraarticular inflammatory cells by 4 hours after injection.39 The mean number of leukocytes present at

4 hrs was 2.4 x 10? cells in the joint wash, and this number continued to increase to a mean of 23.7 ±8.0 cells by 24 hours. Direct comparison with our study is difficult because synovial fluid was obtained by lavage of the joint space with 1 ml of saline, and nucleated cell counts were obtained on the resultant fluid volume. However, these results support our findings of an inflammatory response following intraarticular DIB injection.

We have reported the cytokine data as concentrations and as a relative value because the ELISA kits have not been specifically validated for cytokine analysis in the horse.

However, homology between human and equine II-1 was demonstrated by hybridization of a human II-ip cDNA probe with RNA extracted from E-1 producing equine blood adherent monocytes as determined by RNA immunoblotting. This assay indicates a high degree of homology between the human and the equine proteins. Homology has also been reported between human and murine E-ip, indicating preservation of a functional sequence across species.37 Furthermore, characterization of equine E-1 has been partially done, and showed a high sirrdlarity between the human and equine molecules.^o The EIA used in this study to

163 quantify PGE 2 has been described for use in inflamed synovial fluid in horses.^i Although the manufacturer reports up to 50% cross reactivity to PGEi, PGE 2 has been shown to be preferentially released by equine chondrocytes and synoviocytes in response to a variety of stimuli.42

In conclusion, results of this study illustrate a rapid but mild inflammatory response induced by interleukin-16 in equine joints, that is characterized by increased interleukin 6 and PGE2 release, and a significant neutrophilic vasculitis. Despite short term exposure to this inflammatory cytokine, induction of cartilage matrix degradation was evident. Acute denervation did not influence this inflammatory response, but was protective against the cartilage degradative response. This model could serve as a basis for targeting different inflammatory mechanisms in acute articular inflammation, namely decrease inflammatory cell trafficking, cytokine release and protection of cartilage matrix against an early degradative response.

164 F ootnotes a Cardiomax H, Columbus Instruments, Columbus, OH. b Honeywell VRI2, Electronics for Medicine Inc, Pleasantville, NY c Masterflex variable speed pump. Cole Palmer Int, Chicago, IL d Model 7D Polygraph, Grass instruments, Quincy, MA e Cellular Products Inc., Buffalo, NY f Sherwood medical, St-Louis, MO g Coulter S+IV, Coulter Electronics, Hialeah, FL h Gam Rad, San Juan Capistrano, CA

‘ Advanced Magnetics, Cambridge, MA j TiterZyme PGE2 EIA kit, PerSeptive Diagnostics, Cambridge, MA k Micro-BCA protein assay. Pierce Chemical Co, Rockford, IL

1 Sigma Chemicals, St-Louis, MO m Polyclonal antibody to substance P, Biogenex Laboratories, San Ramon, CA n StrepAvigen Multilink immunodetection System, Biogenex Laboratories, San Ramon, CA

165 Figure 6.1. Illustration of the isolated pump-perfused auto-oxygenated isolated joint model used in the study.

166 Pump Arterial pressure To VR12

FT03 weight a To physiograph Arterial cu’cuit

m m Reference sample Venous pressure

Median artery To physiograph Median vein Synovial fluid

Venous collection Venous circuit

Figure 6.1. Illustration of the isolated pump-perfused auto-oxygenated isolated joint model used in the study.

166 Figure 6.2.Synovial fluid total nucleated cell count across time among groups.

(Group 1 - ■ Control; Group 2 -# Control-Denervated; Group 3- ▲ Inflamed; Group 4-

♦ Inflamed-denervated) ^indicates a significant difference between control joints (Groups

3 and 4) and inflamed joints (Groups 1 and 2) (P=0.001).

167 6 -

330120 180 240 300 330120 Time (minutes)

Figure 6.2

168 Figure 6.3 Synovial fluid total nucleated cell differential (%) over time among groups.

There was a significant increase in % neutrophils and a significant decrease in % large and small monoculear cells across time. There was no difference detected among groups.

169 Neutrophils Large monocytes Small monocytes

Baseline min 300 mm 330 mm240

^ 0.6-

> 0.2 -

Groups

Figure 6.3

170 Figure 6.4. Synovial fluid interleukin IB concentration ratios across time and among groups. *indicates a significant difference compared to baseline (P=0.001). ¥ indicates a significant difference between groups 3 and 4 (P=0.02).

171 Control-innervated

Control-denervated

Inflamed-innervated

V7y Inflamed-denervated

- 35

■ 30 k

- 2 25 - *

- *

- 10 1 - 5- 1 i 0 : - Baseline 240 minutes 300 minutes 330 minutes

Figure 6.4

172 Figure 6.5. Synovial fluid interleukin 6 concentration ratios across time and among groups. *indicates a significant difference between control joints (Groups I and 2) and inflamed joints (Groups 3 and 4) (P=0.01).

173 Control-innervated

Control-denervated

Inflamed-innervated

Inflamed-denervated

- 10

Baseline 240 minutes 300 minutes 330 minutes

Figure 6.5

174 Figure 6.6. PGE2 ratio over baseline across time and among groups. (Group I - ■

Control; Group 2-# Control-Denervated; Group 3- ▲ Inflamed; Group 4- ♦ Inflamed-

Denervated)

* indicates a significant difference in Group 4 (Inflamed-Denervated) (P=0.038).

175 11 Control-innervated ca Control-denervated ■ Inflamed-innervated Inflamed-denervated

8 - 7-

6 5-

4 !3 2

0 240 300 Time (minutes)

Figure 6.6

176 Figure 6.7. Total substance P (pg) in synovial fluid collection from 240-300 minutes

(■) and 300 to 330 minutes (■).* Indicates a significant difference between control joints

(Groups 1 and 2) and inflamed joints (groups 3 and 4) (P=0.025).

177 gg 700

2 500

1 r Control innervated Control denervated Inflamed innervated Inflamed denervated

Figure 6.7

178 Figure 6.8 Synovial membrane histology specimens (lOOX) illustrating A-a control isolated joint from group 1 and B- an inflamed isolated joint from group 3. Inflamed isolated joints (Groups 3 and 4) showed a significant neutrophilic vasculitis (P=0.01).

179 j ' ;

■^mm m \ \ ' .

• v ^

180 Figure 6.9 Synovial membrane substance P and nerve specific enolase immunostaining.

There was no différence among groups as to the number of positively stained fibers.

181 ïp ^

V >

w s*

182 Extent of Edema Hemorrhage 0- None 0- none 1- Less than 25% of intima/subintima 1 - Less than 25% of specimen involved 2- 25-50% of intima/subintima 2- 25-50% of specimen involved 3- 50-75% of intima/subintima 3- 50-75% of specimen involved 4- Greater than 75% of intima/subintima 4- 100% of specimen involved

Neutrophilic vasculitis Type of infiltrate 0- none 1- Vessels only 1- Scattered intraluminal neutrophil clusters, 2- Vesseis/subintima minimal or none intramural or extravascuiar 3- Vesseis/subintima/intima 2-Widespread intraluminal neutrophils, minimal or no intramural 3- < 50% of vessels have intramural clusters; slight perivascular infiltrate oo w 4- > 50% of vessels have intramural clusters; slight perivascular infiltrate 5- Almost ail vessels have intramural and perivascular infiltrates

Table 6.1 Score description for evaluation of synovial membrane histology Concentration Content Ratio Group 1 (Controt-lnnervated) M ean æ Mean æ M ean æ Baseine <1.6 ± 0 .2 5 0 .5 0 ± 0 .1 5 240 minutes <1.6 ±0.43 0.92 ±0.27 300 minutes <1.6 ±0.29 3.25 ±0.65 0 .6 9 ± 0 .1 8 330 minutes <1.6 ± 0 .3 4 2.71 ±0.61 0.75 ±0.21 Total content 5.96 ± 1 .2 4 Group 2 (Control-Deneivated) Base#ne <1.6 ±0.17 0.45 ±0.21 240 minutes <1.6 ±0.16 0.21 ±0.11 300 minutes <1.6 ±0.16 1.06 ± 0.48 0.31 ± 0 .1 5 330 minutes <1.6 ±0.18 0.54 ±0.20 0.42 ±0.15 Total content 1.59 ±0.58 Group 3 (Inflamed-lnnecvated) Baseine <1.6 ±0.36 1.10 ±0.46 240 minutes 28.23 ±13.26*11 19.07 ±8.96*¥ 300 minutes 16.99 ±7.25*¥ 120.24 ±37.26*¥ 15.44 ±6.58*¥ 330 minutes 10.67 ±5.10*¥ 44.36 ±15.11*¥ 8.89 ±4.25*¥ Total content 164.60 ± 4 6 .7 1*¥ Group 4 (Inflamed-Denervated) Baseine <1.6 ±0.09 0 .4 6 ±0.11 240 minutes 12.74 ±3.43* 8.61 ±2.31* 300 minutes 3.65 ±0.75* 75.05 ±18.23* 3 .3 2 ±0.68* 330 minutes 2 .3 7 ±0.61* 32.14 ±12.34* 1.97 ±0.51* Total content 107.19 ±28.18*

Table 6 2 Synovial fluid 016 concentration, content and ratio among groups and across time. Ratio calculated using Group 1 as reference. * indicates inflamed joints (Groups 3 and 4) significandy different from controls (Group 1 and 2) (P<0.(XX)1) ¥ indicates inflamed irmervated joints (group 3) signifrcandy greater than inflamed-denervated joints(Group 4) (P=0.04)

1 8 4 Concentration Content Ratio Group 1 (Control-lnnervaterfl Mean SE Mean æ Baseline < 3.2 ±0.00 0.00 ±0.00 240 minutes <3.2 ±0.38 0.32 ±0.12 300 minutes 5.68 ±2.64 7.78 ±2.61 1.78 ±0.82 330 minutes 6.18 ±2.17 13.28 ±4.26 1.93 ±0.68 Total content 21.05 ± 6 .8 0 Group 2 (Control Denervated) Basefine <3.2 ±0.01 0.00 ±0.00 240 minutes <3.2 ±0.84 0.33 ±0.26 300 minutes 3 .3 0 ±2.81 7.02 ± 5 .8 4 1.03 ±0.88 330 minutes 2 .2 3 ±1.79 0.96 ±0.41 0.70 ±0.56 Total content 7.98 ± 5 .6 6 Group 3 (Infiamed-lnnervated) Basefine <3.2 ±0.20 0.08 ±0.06 240 minutes 28.46 ±23.88* 8.89* ±7.46 300 minutes 4 3 .4 7 ±34.65* 213.15 ±108.81* 13.58* ±10.83 330 minutes 43.73 ±31.61* 250.69 ±149.5* 13.66* ±9.88 Total content 463.84 ±256.15* Group 4 (Inflamed-Denervated) Baseline < 3 .2 ± 0.43 0.1 7 ±0.14 240 minutes 46.01 ±25.71* 14.38* ±8.04 300 minutes 59.81 ±30.52* 551.66 ±329.02* 18.69* ±9.54 330 minutes 68.79 ±33.60* 584.89 ±357.27* 21.49* ±10.50 Total content 1136.56 ±684.78*

Table 6.3 Synovial fluid 116 concentration, content and ratio among groups and across time. Ratio calculated using Group 1 as reference. * indicates inflamed joints (Groups 3 and 4) significantly different from controls (Group 1 and 2) (P=0.01)

185 Proteoglycan aynthaala 18 hrs 72 hrs 144 hrs Control Isolated Ratio Control Isolated Ratio Control Isolated Ratio

Group 1 (Control Innervated) 8.14 ±2.32 7.49 ±2.28 0.97 ±0.16 17.52 ±5.65 19.36 ±6.32 1.09 ±0.13 15.73 ±7.71 12.97 ±4.94 0 .98 ±0.17 Group 2 (Control Denervated) 7.38 ±1.43 6.47 ±1.63 1.00 ±0.23 23.66 ±12.59 21.19 ±6.12 1.41 ± 0.44 12.47 ±3.39 15.49 ±6.07 1.16 ±0.38 Group 3 (Inflamed-lnnetvated) 23.78 ±8.98 23.68 ±7.06 1.21 ±0.20 80.26 ±52.82 82.16 ±45.18 1.39 ±0.44 46.89 ±28.12 33.07 ±14.13 0 .98 ±0.21 Group 4 (Inflamed Denen/ated) 10.74 ±2.31 13.18 ±3.45 1.20 ±0.18 27.49 ±7.66 44.60 ±17.48 1.50 ±0.24 19.01 ±5.40 23.03 ±8.48 1.17 ±0.30

Protaoglycan content 18 hrs 72 hrs 144 hrs Control Isolated Ratio Control Isolated Ratio Control Isolated Ratio Group 1 (Control Innervated) 0.20 ±0.02 0.22 ±0.03 1.14 ±0.14 0.1 8 ±0.04 0.2 3 ± 0.08 1.24 ±0.16 0.15 ±0.02 0.1 7 ±0.01 1.14 ±0.07 ^ ro u p 2 (Control-Denervated) 0.24 ±0.04 0.19 ±0.04 0.87 ±0.15 0.1 3 ±0.04 0.1 3 ± 0.04 1.03 ±0.12 0.12 ±0.03 0 .1 0 ±0.03 0.78 ±0.22 ° Group 3 (Infiamed-lnnervated) 0.31 ±0.04 0.28 ±0.04 0.93 ±0.13 0.1 8 ±0.03 0.1 9 ± 0.03 1.21 ±0.29 0.14 ±0.04 0 .1 9 ±0.03 1.54 ±0.29 Group 4 (Inflamed-Denervated) 0.27 ±0.04 0 .2 3 ±0.04 0.93 ±0.09 0.1 8 ±0.02 0 .1 6 ± 0.03 1.04 ±0.22 0.1 2 ± 0.02 0.14 ±0.03 1.12 ±0.16

Table 6.4 Proteoglycan synthesis (CPM/|xg protein) and content (pg/pg protein) and respective ratio of isolated to contralateral control joint among groups and at 18 hours, 72 hours and 144 hours time in culture. There was no significant difference among groups or across time. Group 1 1 Group 2 1 Group 3 Group 4 (Control innervated) (Control Denervated) (Inflamed Innervated) (Inllamed Denervated) Total glycosaminoglycan 18 hrs Control 60.26 ±12.76 101.03 ±19.62 132.72 ±34.69 73.50 ±10.12 Isolated 57.07 ±8.44 82.21 ±12.06 119.37 ±25.63 92.75 ±11.98 Ratio 1.20 ±0.33 0.67 ±0.07 1.02 ±0.16 1.34 ±0.15 72 hrs Control 22.13 ±6.98 34.41 ±12.34 34.11 ±6.97 33,70 ±4.45 Isolated 17.51 ±3.46 36.61 ±13.99 43.37 ±10.48 45.22 ±10.07 Ratio 1.14 ±0.29 1.23 ±0.28 1.35 ±0.24 1.49 ±0.42 144 hrs Control 45.21 ±10.47 63.03 ±14.81 81.14 ±18.28 34.50 ±6.79 Isolated 48.49 ±11.79 45.73 ±12.05 58.97 ±17.20 45,01 ±9.92 Ratio 1.13 ±0.16 0.73 ±0.13 1.05 ±0.22 1.18 ±0.19 Chondroitin sulfate 18 hrs Control 36.66 ±11.56 68.79 ±12.77 103.46 ±30.19 53.44 ±9.62 Isolated 35.06 ±9.33 55.71 ±6.96 69.71 ±24.65 63.21 ±7.23 Ratio 1.13 ±0.50 0.88 ±0.09 0.97 ±0.19 1.60 ±0.44 72 hrs Control 7.15 ±2.02 23.25 ±8.79 22.18 ±7.37 18.54 ±3.79 23 isolated 7.20 ±3.07 23.20 ±9.79 28.02 ±11.69 31.06 ±7.29 Ratio 0.73 ±0.30 1.21 ±0.38 2.18 ±1.04 2.24 ±0.73 144 hrs Control 24.59 ±4.50 39.12 ±11.03 43.68 ±13.01 17.32 ±5.97 Isolated 29.63 ±6.91 26.09 ±10.31 36.19 ±14.57 32.66 ±8.17 Ratio 1.26 ±0.35 0.64 ±0.14 0.75 ±0.26 6.97 ±7.76 Keratan sulfate 18 hrs Control 9.50 ±5.00 7.80 ±1.97 13.80 ±4.98 6.33 ±3.25 isolated 11.58 ±4.34 8.71 ±3.91 14.19 ±5.30 14.20 ±6.88 Ratio 1.54 ±0.63 0.96 ±0.17 0.78 ±0.26 3.57 ±1.57 72 hrs Control 9.49 ±6.01 2.96 ±1.98 10.64 ±4.60 10.80 ±5.29 Isolated 6.11 ±2.91 6.76 ±3.14 10.59 ±5.18 7.69 ±4.53 ratio 2.04 ±1.72 1.96 ±0.76 0.61 ±0.31 0.62 ±0.22 144 hrs Control 8.04 ±2.87 10.23 ±4.43 13.25 ±6.19 10.70 ±6.28 Isolated 7.89 ±5.67 8.99 ±5.57 17.59 ±7.52 4.23 ±1.98 Ratio 0.71 ±0.48 0.54 ±0.20 0.99 ±0.34 0.72 ±0.44

Table 6.S Cartilage total glycosaminoglycan, chondroitin and keratan sulfate concentrations among groups at culture time 18,72 and 144 hours. There was no significant difference among groups or across time. Degradation of newly synthesized proteoglycan Group 1 Group 2 Group 3 Group 4 (Control-Inneryated) (Control-Deneryated) (Inllamed Inneryated) (Inflamed-Deneryated) 24 hrs Mean SE Mean SE Mean SE Mean SE Amount released (CPM) Control 72261 2 1 0 8 8 7 5 5 8 7 2 9 4 8 0 179223.4 81283.54 74578.14 189 8 9 .3 8 Isolated 8 4 1 2 9 35207 73173 29735 185019.8 81550.46 5 5 1 4 2 .5 4 14802.27 Ratio 1.02 0.28 1.34 0.50 0 .9 0 0.12 0.71 0.07" % released Control 0 .4 0 0 .08 0.41 0 .07 0.39 0.05 0.35 0.04 Isolated 0 .4 0 0 .07 0 .3 9 0.08 0.33 0.08 0.32 0.05 Ratio % released 1.50 1.07 1.00 0.13 0.90 0.13 0.92 0.08 48 hrs Amount released (CPM) Control 28737 8970 38248.59 21252.09 80918.5 14282.55 49837.91 15993.97 Isolated 3 5 5 4 9 17774 82381.21 27430.83 77663.3 14230.37 41510.08 13005.38 Ratio 0.92 0.12 1.90 0.88 1.40 0.29 0.8 4 0.07" % relased Control 0.15 0.02 0.18 0 .05 0.14 0.02 0.21 0.03 2S Isolated 0.18 0.02 0 .2 5 0.08 0.23 0.08 0 .22 0 .0 3 Ratio % released 1.13 0.14 1.80 0.45 1.59 0.45 1.09 0.04 72 hrs Amount released (CPM) Control 22857 7420 18888 4 5 2 9 4 3 8 8 8 11593 3 1 9 0 7 11085 Isolated 2 5 8 8 8 10275 28841 9581 41089 9758 30349 11972 Ratio 1.14 0.24 1.27 0.35 1.00 0.19 0.92 0 .09 % relased Control 0.14 0.0 4 0.12 0.03 0.10 0.02 0 .13 0.01 Isolated 0 .1 8 0.04 0.13 0.03 0.0 9 0.01 0 .1 6 0.0 2 Ratio % released 1.24 0.2 2 1.07 0.14 1.00 0.16 1.19 0 .08

Table 6.6 Newly synthesized proteoglycan released in 24,48 and 72 hr media among groups expressed as amount released (CPM), ratio of isolated to contralateral control joints, and % of total cartilage content. ♦ indicates a significant decrease in release in inflamed denervated Joints (P=0.03) 188 Degradatlon of endogenous proteoglycan Group 1 Group 2 Group 3 Group 4 (Control-Innervated) (Control-Denervated) (Infiamed-lnnervated) (Inflamed-Denervated) 24 hrs Mean SE Mean EE Mean SE Mean SE Amount released (p 3) Control 46.19 2.17 50.78 7.03 62.86 12.93 59.61 3.04 Isolated 48.73 4.61 58.94 12.26 56.04 13.80 48.54 4.07 Ratio 1.07 0.12 1.15 0.11 0 .8 8 0 .13 0.81 0 .06 % relased Control 0 .2 2 0 .03 0 .2 5 0.0 2 0.31 0 .05 0 .2 8 0 .02 Isolated 0.21 0 .02 0 .2 7 0.01 0 .2 3 0.07 0 .2 5 0 .03 Ratio % released 1.02 0.11 1.10 0.0 7 0 .7 9 0 .22 0.91 0.11 48 hrs Amount released (p 1) Control 50.94 1.78 52.86 5.42 49.31 7.78 58.27 6.45 Isolated 60.86 1.35 69.30 15.64 54.55 12.30 70.25 8.11 Ratio 1.20 0 .05 1 .26 0 .1 6 1.05 0 .19 1.21 0 .07 % relased Control 44.05 9.86 0 .2 7 0.02 0 .2 4 0 .04 0 .2 6 0.01 Isolated 57.75 17.22 0 .3 2 0.0 2 0 .1 8 0,07 0 .3 5 0 .0 3 Ratio % released 1.13 0.1 0 1.19 0 .06 0.71 0 .18 1.32 0 .06 72 hrs Amount released (p 1) Control 58.90 10.07 53.71 5.04 53.41 6.53 60.06 7.42 Isolated 50.74 2.16 48.62 7.02 52.65 9.93 47.11 7.19 Ratio 0 .9 4 0 .12 0 .8 9 0 .0 9 0 .9 8 0,11 0 .8 4 0 .1 3 % relased Control 0 .2 6 0 .02 0 .2 7 0.01 0 .1 9 0 ,05 0 .2 7 0 .02 Isolated 0 .22 0.02 0 .2 4 0.0 4 0 .3 3 0,17 0 .2 3 0 .0 3 Ratio % released 0 .9 0 0.12 0 .8 7 0.11 1.62 0.64 0 .8 9 0 .12

Table 6.7 Endogenous proteoglycan released in 24,48 and 72 hr media among groups expressed as amount released (pg) ratio of isolated to contralateral control joints, and % of total cartilage content. There was no difference among groups or across time. Edema Hemorrhage Neutrophilic vasculitis Type of Infiltrate Control Isolated Control Isolated Control Isolated Control Isolated

Fibrous synovium Group 1 (Control-Innervated) 0.1 7 ±0.11 1.22 ±0.55 0.00 ±0.00 0.00 ±0.00 0.4 4 ±0.20 1.78 ±0.67© 0 .4 4 ±0.20 1.11 ±0.36 Group 2 (Control-Denervated) 1.47 ± 0.62 1 .39 ±0.62 0.1 3 ± 0 .1 3 0.17 ± 0.17 0.33 ±0.11 1.50 ±0.51© 0.33 ±0.11 1.00 ±0.31 Group 3 (Inflamed Innervated) 1.50 ± 0.62 0.56 ±0.20 0.00 ±0.00 0.00 ±0.00 0.28 ±0.06 2.22 ± 0 .4 8 © t 0 .2 8 ±0.06 1.44 ±0.27 Group 4 (Inftamed-Denenrated) 0.38 ±0.29 0.90 ±0.40 0.00 ± 0.00 0 .29 ± 0.18 0.2 4 ± 0.06 2.81 ±0.51 © t 0 .2 4 ±0.06 1.81 ±0.32

Villous synovium Group 1 (Control-lnnen/ated) 1.06 ± 0,47 0.93 ±0.46 0.00 ±0.00 0.13 ±0.13 0.67 ± 0 .1 7 ¥ 2 .1 3 ±0.48¥© 0.67 ±0.17 1.33 ± 0.33 Group 2 (Control-Denervated) 0.80 ±0.20 0 .7 2 ±0.35 0.00 ±0.00 0.28 ±0.28 0.40 ± 0 .1 2 ¥ 1 .83 ± 0 .67¥© 0 .4 0 ±0.12 1.00 ±0.30 Group 3 (Infiamed-lnnervated) 1.28 ±0.29 1.44 ±0.43 0.0 0 ±0.00 0.44 ± 0 .3 3 0 .20 ± 0 .1 3¥ 2 .8 3 ± 0 .4 0 ¥ © t 0 .2 0 ± 0.13 2.06 ±0.35 s Group 4 (Inftamed-Denenrated) 0 .6 7 ± 0.36 1.3 3 ±0.29 0.05 ±0.05 0.38 ±0.28 0.67 ± 0 .3 4 ¥ 3 .8 6 ± 0 .2 6 ¥ © t 0 .5 2 ±0.20 2.3 3 ±0.22

Table 6.8 Histopathology scores (mean ± SE) among groups, sites, and comparing contralateral control to isolated joints. ¥ indicates significant difference between villous and fibrous synovium. © indicates significant difference between isolated (Groups 1-4) and contralateral control Joints, t indicates significant difference between Groups 3 and 4 and Groups 1 and 2 References

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29. Miao, F. J.-P., W. Janig, J. D. Levine. Role of sympathetic postganglionic neurons in synovial plasma extravasation induced by bradykinin. J Neurophysiology 75:715-724, 1996.

192 30. Gueme, P.-A., B. L. Zuraw, J. H. Vaughan, et al. Synovium as a source of interleukin 6 in vitro. J Clin Invest 83:582-592, 1989.

31. Shinmei, M., K. Masuda, T. Kikuchi, et al. The role of cytokines in chondrocyte mediated cartilage degradation. J Rheumat Suppl 18:32-34,1989.

32. Blackburn, W. D. J., L. Heck, L. D. Loose. Inhibitions of 5-lipoxygenase product formation and polymorphonuclear cell degranulation by tenidap sodium in patients with rheumatoid artimtis. Arhtritis Rheum 34:204-210,1991.

33. Tammanini, C., E. Seren, G. Pezzoli. Prostaglandin E1-E2 concentration in synovial fluid of horses affected with arthropathy. Clin Vet 103:544-549, 1980.

34. Fairbum, K., M. Kunaver, L. S. Wilkinson, et al. Intercellular adhesion molecules in normal synovium. Brit J Rheumatol 32:302-306, 1993.

35. Morris, E. A., B. V. Treadwell. Effect of interleukin 1 on articular cartilage from young and aged horses and comparison with metabolism of osteoarthritic cartilage. Am J Vet Res 55:138-146, 1994.

36. Alwan, W. H., S. D. Carter, J. B. Dixon, et al. Interleukin-1-like activity in synovial fluids and sera of horses with arthritis. Res Vet Sci 51:72-77, 1991.

37. Morris, E. A., B. S. McDonald, A. C. Webb, et al. Identification of interleukin-1 in equine osteoarthritic joint effusions. Am J Vet Res 51:59-64, 1990.

38. May, S. A., R. E. Hooke, P. Lees. Species restriction demonstrated by the stimulation of equine cells with recombinant human interleukin-1. Vet Immunol Immunopath 30:373-384, 1992.

39. Henderson, B., E. R. Pettipher. Comparison of the in vivo inflammatory activities after intraarticular injection of natural and recombinant IL-1 alpha and EL-1 beta in the rabbit. BiochemPharm 37:4171-4176, 1988.

40. May, S. A., R. E. Hooke, P. Lees. The characterisation of equine interleukin-1. Vet Immunol Immunopath 24:169-175, 1990.

41. Owens, J. G., S. A. Barker, S. A. Stanton. Measurement of prostaglandin E2 (PGE2) and leukotriene B4 (L1B4) using an enzyme-linked immunosorbent assay (ELISA) in horse plasma and syovial fluid. Toxicologist 13:147, 1993.

42. May, S. A., R. E. Hooke, P. Lees. Identity of the E-series prostaglandin produced by equine chondrocytes and synovial cells in response to a variety of stimuli. Res Vet Sci 46:54-57, 1989.

193 SUMMARY

Articular inflammation is a common finding in equine joints disorders. As in other organs, several aspects of the joint system, including blood flow, transsynovial permeability and fluid exchanges, cellular infiltration and mediator release may be involved in the inflammatory process and contribute to joint morbidity. With inflammation, tissues within the joint contribute to mediator release and stimulate degradation of the cartilage matrix. Perfused oxygenated extracorporeal isolated models have been thoroughly described for organs such as heart, lung and intestine, but have never been created for the joint organ. The studies reported here describe an isolated pump-perfused auto-oxygenated extracorporeal isolated joint model developed in the equine. Furthermore, these studies characterize local Starling’s forces in the isolated joint at isogravimetric state and alterations in Starling’s forces in response to hemodynamic manipulations. Subsequently, these studies report on local hemodynamic, fluid exchanges, metabolic responses, alterations in permeability to small and large molecules, inflammatory mediator release, cellular events and articular cartilage metabolic alterations in experimentally induced acute articular inflammation in the isolated, and furthermore innervated or denervated, joint.

194 decreased exponentially. This study also revealed an important function of synovial fluid to increase joint compliance and hysteresis therefore preventing collapse of the joint cavity and allow fluid exchanges at subatmospheric pressures. Furthermore this study emphasized the importance of joint angle on pressure-volume relationships. In the second preliminary study, a colored microsphere technique was validated for determination of blood flow to the synovial membrane and joint capsule. Furthermore, that study documented a significant decrease in synovial membrane blood flow at intraarticular pressures of 30 mmHg or greater.

The development and physiologic responses of the isolated model were subsequently described. The extracorporeal pump perfused isolated joint unit developed in this model was successfully be maintained for 6 hours. Using this system, isogravimetric state was defined at a circuit arterial pressure of 133 mmHg and venous pressure of 10.5 mm Hg. Starling’s forces were defined for this system. Arterial and venous pressure manipulations revealed that increases in arterial pressure significantly raised transsynovial flow and synovial fluid production, when LAP was maintained at atmospheric pressure. A similar but much less important response was observed with increased venous pressure.

Acute inflammation was successfully induced in the isolated innervated or denervated joint preparation, using interleukin-IB. Acute inflammation induced a significant increase in oxygen consumption and extraction ratio in response to increased metaboUc demand, acute inflammation also increased synovial membrane permeability to albumin, while innervation was associated with a higher permeability to large molecules (MW

144,(X)0). Inflammation also significantly increased synovial fluid production and the permeability surface area product of the synovial membrane

195 detected in synovial fluid collected from inflamed joints. Inflammation induced a significant degradative response in articular cartilage, even after this short term exposure to an inflammatory mediator.

196 APPENDIX A DYNAMIC VASCULAR AND TRANS-SYNOVIAL FORCES

OF THE ISOLATED JOINT

Introduction

It has been widely presumed that Starling’s forces govern trans-synoviai fluid flow as in other connective tissue spaces.29.54 Starling’s hypothesis predicts that the rate of transcapillary fluid filtration (Jy) is a linear function of capillary pressure (Pcap) and interstitial colloid osmotic pressure and a negative linear function of plasma colloid osmotic pressure and interstitial pressure (PJ. Due to the presence of a large third space (joint cavity), a simple two-compartment model of fluid exchange with the interstitium does not necessarily apply. Many sophisticated and elaborate studies have been performed to indirectly measure or theoretically calculate the primary coefficients modifying the fluid flow from plasma to synovial fluid (osmotic coefficient (C7d) and filtration coefficients of the capillary (Kc) and synovium (Xs)).36 An isolated, rabbit perfused hindlimb preparation has been used to describe trans-synovial flow, but pressure changes and blood flow were not representative of only the joint and trans-synovial flow was in the absorptive p h a se.2 9

Presumption and theoretical calculation of hemodynamic, microvascular, and trans- synovial parameters, therefore, were required due to a lack of an isolated joint model.

Isolated organ preparations have facilited these measurements in the heart, lung, intestine, and hoof laminae.2.3.30,6i An isolated joint model would more accurately permit simultaneous measurement of dependent changes in blood flow, synovial fluid production

197 and composition, and interstitial fluid accumulation as expected to occur with physiologic manipulation, or pathology such as acute joint inflammation or degenerative joint disease J

Few studies have evaluated physiologic parameters of the joint separately, includinglymphatic flow .47 Clearance studies from the synovial cavity following intraarticular injection of i33Xenoni4 or N a i2 3 l and 13I-I labeled album in® ) can estimate blood flow, but are limited by radioactivity, diffusion, and the inflammatory response. It has been demonstrated that an intra-articular injection of even balanced electrolyte solution produces transient, but significant, synovitis that alters blood flow .58 Metabolic parameters in synovial fluid (pCOg, pH, and lactate concentration) have been measured to indirectly estimate blood flow relative to oxygen demand, and estimate joint ischemia, but they do not provide direct and simultaneous measurements of fluid exchange.27 In virtually all studies, measured values of filtration or trans-synovial fluid flow were either theoretically calculated, inherently inaccurate, or limited by the number of physiologic parameters that could be evaluated. Dynamic evaluations of the vascular/fluid component(s) of joints is critical to understanding joint physiology, pathophysiology and anticipation of responses to medications. Therefore, the purpose of this study was to develop an autooxygenated, pump-perfused, isolated joint model and use this model to determine the hemodynamic and trans-synovial forces and characteristic responses of joint tissue to controlled physiologic (pressure) manipulations.

The horse was selected for use in an isolated joint preparation due to anatomical considerations, including a well-developed singular digit (Mem) with a simple neurovascular network and lack of musculature. The joint selected (metacarpophalangeal

[MCP] joint ) is large enough to allow collection of ample quantities of synovium and synovial fluid for multiple analyses. Also, the MCP joint is the second most common joint affected with arthritis in horses and therefore would have future clinical relevance to the

198 study of naturally-occurring arthritis. Equine osteoarthritis is naturally-occurring, histologically similar to human osteoarthritis and has been extensively studied due to the impact of osteoarthritis on the equine racing industry. Many studies of equine osteoarthritic cartilage (natural or induced) have confirmed similar behavior of equine chondrocytes and articular tissues as in man and other species.4.6.12. ig. 19 .40.4 1.44 .45 .48 .49 .57,62 Substance P innervation and receptor localization in the equine joint are similar to man.9 Comparatively in the horse, joint disorders are also the most common cause of lameness and loss of athletic use.28.38

Materials and methods

Horses

Seven normal horses (aged 1-5 years [mature, but young]; 306-556 kg body weight; Thoroughbred-type), normal as determined by physical and lameness examination,

CBC and serum chemistry profile, were obtained. Inclusion criteria for the study dictated that both MCP joints were palpably normal with a normal range of motion (~ 90° ), normal radiographic evaluation (4 standard views), and normal synovial fluid analysis. Gross evaluation of articular cartilage at the termination of the study had to be normal, or the horse was eliminated from the study and replaced. All experiments were conducted in accordance with the Animal Care and Use Guidelines of The Ohio State University.

Anesthesia and Systemic Hemodynamic Measurements

Only one horse was studied on a given day because of the complexity of the preparation. Horses were sedated with xylazine (0.5 mg/kg, iv), and anesthesia was induced with guaifenisin (-25 mg/kg) followed by sodium thiopental (2 mg/kg, iv).

Anesthesia was maintained with sodium pentobarbital (5-15 mg/kg/hr, iv, to effect) and

199 controlled mechanical ventilation with 100% oxygen. Horses were positioned in lateral recumbency, and anesthesia monitored by continuous display of direct systemic arterial blood pressure measurement, hourly measurement of cardiac output by a thermodilution technique, and intermittent arterial pH and blood gas measurements. Mean systemic arterial blood pressure was maintained at > 70 nun Hg, and ventilation was adjusted to maintain

PaC02 < 55 mmHg. Heparin (50,000 units IV) was administered every 90 minutes.

Joint isolation procedure and Instrumentation

An MCP joint of one forelimb was randomly selected. Before joint isolation, an arterial cannula (PE 240) was placed in the median artery proximal to the anticipated site of joint isolation. Blood was diverted from the cannulated median artery, circulated through a calibrated peristaltic pumpa, and infused via the cannulated medial palmar artery to the isolated joint preparation. (Fig 1) Venous outflow was collected from the medial palmar and lateral and medial palmar digital veins, directed into a reservoir and reinfused into the horse via the median vein using a peristaltic pumpb. Weight of the preparation was continuously measured using an FT03 transducer^ connected to a physiography for continuous display. Circuit arterial pressure (Pacir) was measuredeJ and initially set at 150 mmHg by adjustment of flow in the arterial pump, and circuit venous pressure (Pvcir) was adjusted using a clamp and initially maintained at 10 mmHg. Pressure tranducers were placed at the level of the joint preparation. Final isolation of the MCP joint was achieved by transecting all soft tissue and sectioning bone at the level of the distal metacarpus and dislocation of the proximal interphalangeal joint. (Fig 1). A distended lymphatic vessel visible around the catheterized lateral palmar artery was carmulated with a fine 26 g catheter for collection of lymph. The catheter was capped between collections. A 22 g intraarticular catheter was placed in the dorsal pouch of the MCP joint and attached to a shielded

200 photoelectric cell drop counter. The drop counter was connected to the physiography for continuous display. The synovial fluid was continuously drained and collected to maintain atmospheric intraarticular pressure (lAP) and determine trans-synovial fluid flow (Qs). In this system, 16.5 drops was approximately 1 ml of fluid. Synovial tissue blood flow was measured by the colored microsphere technique at three separate times; before joint isolation, after 15 minutes of equilibration in the isogravimetric state (baseline) and after

Pacir manipulations and return to an isogravimetric state. Body temperature of the joint preparation was maintained with a heat lamp.

Colored Microsphere Technique

Synovial blood flow in the isolated joint preparation was measured at three time points in the experiment using a colored microsphere technique.22.32.43 Blue, red or yellow colored microspheress were injected into the arterial inflow line and a reference arterial blood sample was withdrawn from the medial palmar digital artery at a rate of 4.94 ml/min for 1 minute. (Fig 2) Tissue specimens and reference blood samples were digested in KOH and filtered through an 8 |im pore size polyester filter. Dimethylformamide was added to release the dye from the microspheres. Dye concentration was measured by spectrophotometry using a matrix inversion technique for resolution of composite spectra when multiple colors are used. Blood flow was expressed as ml/min/g of tissue.29.33

Baseline Vascular Measurements and Calculations

The isogravimetric state (no gain or loss of weight in the isolated preparation) was maintained for 25 minutes before collecting baseline data (5 minutes); circuit arterial blood flow (2acir;ml/min), venous blood flow (j2vcir;nil/min), lymphatic flow (g/;ul/min), synovial flow (2s;drops/min), and interstitial fluid accumulation (preparation weight).

201 Synovial fluid was collected during this 5 minute baseline period as a representative sample for the previous 30 min manipulation and flow expressed as number of drops per minute over this 5 minutes. Capillary pressure (Pcap) was determined using the venous occlusion technique during the isogravimetric state, and pre- and postcapillary resistance and pre- and post capillary resistance ratios were calculated.^ Total vascular resistance, precapillary resistance and postcapillary resistance were determined by dividing the arterial-venous, arterial-capillary, and capillary-venous pressure gradients, respectively, by blood flow.2

Lymph, plasma and synovial fluid was collected and placed in a capillary tube for total protein and albumin determination and calculation of oncotic pressure, osmotic reflection coefficient and transitional microvascular pressures.

Calculations were made for total vascular resistance (/?t) of the isolated joint preparation (Rt = Pacir - Pvdr / Q ^r)l vascular compliance (% change in preparation weight during the immediate elevation in the vascular phase of the weight gain curve/ change in pressure (assuming the filtrate has a unit density and that the change in pressure equals the difference between two sequential Pacir or Pv^r), and tissue compliance (% change in preparation weight during the last 5 minutes of each pressure manipulation trial / change in pressure).2

Measurement o f synovial fluid, plasma and lymph total protein concentration

Total protein concentration was determined using a Dye binding assay*» using bovine serum albumin as a standard. Synovial, semm and lymph albumin fractions were measured by electrophoresis. Protein (total and albumin) was used for calculation estimates of oncotic pressures for plasma synovial fluid {its), and lymph (/tQ; osmotic reflection coefficient (CTd); net filtration pressure; and transitional microvascular pressure

202 (77lfP).2.18,21,53^6 Fluid oncotic pressure {k) was estimated by the calculation formula k

= 1.4C + 0.22C2 + 0.005 C3, where C is the concentration of the protein in the fluid.2

Fluid osmotic reflection coefficient was calculated as I-Cj/ Cp where Q is the concentration (g/lOOml) of protein in the synovial fluid and Cp is the concentration of protein in the plasma collected at the same time period. Calculation was made using the steady-state relationship between synovial fluid and plasma protein concentration ratio

(Cs/Cp) and synovial fluid flow in a similar fashion as has been done with lymph flow.2

Synovial fluid flow was increased with by increasing Pv^r until C^/Cp reached a steady- state, filtration-independent phase.3.i8 The was estimated by using 1-Cg/Cp, where

Cs/Cp was nitration-independent.2.3,18 Net filtration pressure was calculated as the difference between the outward and inward capillary forces in the Starling equation [net filtration pressure = (Pcap + - (Pi + ;i|,)].2i Interstitial fluid pressure (P,) was zero because lAP was zero and the equation was reduced to [net filtration pressure = Pcap + ^ -

Jt^. The calculation of net filtration pressure was modified as follows to account for tissue characteristics since Oj could be calculated in our MCP preparation [net filtration pressure

= (Pcap - Pi) - Od ( ^ — 18 ,20,21 Transitional microvascular pressure for synovial fluid flow (TMPs) was calculated using the formula TM P^ 7Cp- 7Cs + Ps where is the oncotic pressure in plasma, is the oncotic pressure in synovial fluid and P$ is the hydrostatic pressure of the synovial fluid.53 Hydrostatic synovial fluid pressure was zero in the preparation since synovial fluid was removed and maintained at atmospheric pressure, therefore the formula was reduced to TMPs = it^-7ts- Transitional microvascular

203 pressure estimates the theoretical pressure at which filtration and absorption forces are in balance. A TMP for lymph flow (TMPi) was estimated similarly. Lymph hydrostatic pressure was zero and lymph flow was zero.

Pressure Manipulations

After baseline measurements including tissue blood flow (microspheres) and establishment of the isogravimetric state, Pa^r was adjusted to 100, 150,200 and 250 mmHg for 30 minutes and circuit arterial pressure (Pa^r), circuit arterial (QScir) and venous

(<2vdr) flow, cardiac output (CO), systemic arterial pressure (f^; systolic and mean), synovial fluid production (Qs), lymph production (Qi) and weight changes (vascular and tissue compliance) were recorded during the last 5 minutes of the 30 minute period at each pressure. Pa^r was returned to the isogravimetric state for 30 minutes before venous pressure manipulations (Pvcir). Tissue blood flow (microsphere) and capillary pressure

(Pcap) were recorded during the last 5 minutes of the isogravimetric condition. Capillary pressure was measured using the venous occlusion method. i P v ^ r was adjusted to 2 0 and

40 mmHg and measurements made as described above. Total time of the isolated joint preparation was a mean of 3.5 hours. The total anesthesia time was a mean of 4.5 hours.

The MCP joint was injected with Evans blue-albumin for gross inspection of the articular cartilage after euthanasia and recording the above data.

Gross evaluation of the joint

Horses were humanely euthanized with an overdose of sodium pentobarbital (iv).

The MCP joint was opened and the cartilage visually inspected for any evidence of osteoarthritic cartilage (surface fibrillation, erosion or fraying). The Evans-blue dye

204 enhanced the visibility of these lesions. Horses with any evidence of osteoarthritis were eliminated from the study.

Synovium was harvested for microsphere analysis immediately after inspection of the cartilage. The joint capsule and synovial lining were collected from the dorsal and palmar joint pouches. The synovial membrane was dissected from the fibrous layer of the joint capsule and separated into 0.5-2.0 g specimens that were processed as described for microsphere analysis.

Statistical Analysis

A non-parametric repeated-measures (time) analysis (Friedman test) was used to determine differences among the variables measured for each pressure manipulation. A

LSD post-test specifically located the differences among pressure manipulations.

Significance was set at p< 0.05.

Results

Data reported are as the mean +/- SEM from horses (n=7) that met all the inclusion criteria. (Table 1) No horses were eliminated from the study.

Anesthesia and Systemic Hemodynamics

All horses met the anesthesia inclusion criteria (mean systemic arterial blood pressure > 70 mm Hg, >150 mm Hg and PaCOi < 55 mmHg) for the duration of the study. (Table 2) Mean systemic arterial pressure ranged from 106-136 mmHg and systolic arterial pressure ranged from 142-164 mmHg and did not significantly change during the

4.5 hours of study (p = 0.56 and 0.84, respectively). Cardiac output significantly

(p=0.005) decreased from 0.077 L/min/kg at the beginning of anesthesia to 0.045 L/min/kg

205 at the end of the study. Arterial oxygen tension (PaC> 2) was highest at the beginning of anesthesia, lowest at the end of the anesthesia and ranged from 359.5-461.3 mmHg

(p=0.05) . Correspondingly, arterial carbon dioxide tension (PaCOi) ranged from 37.4-

54.8 mmHg and significantly increased with the duration of anesthesia (p=0.01).

Tissue Perfusion

Synovial membrane blood flow before joint isolation was a mean of 0.073 ml/min/g and did not significantly change at isogravimetric conditions immediately after joint isolation (mean 0.16 ml/min/g) or at isogravimetric conditions near the termination of the study (mean 0.049 ml/min/g) (p=0.26). (Table 2) The palmar sites tended to have greater tissue blood flow (mean 0.261 ml/min/g) than the dorsal sites (mean 0.058 ml/min/g) only immediately after isolation (p=0.083).

Baseline and Isogravimetric Characteristics

Total blood flow and arterial pressure in the palmar artery before joint isolation

(i.e., to the distal limb) was 68 ml/min and 149 mmHg, respectively. Venous pressure taken from the distal palmar vein was 20.0 mHg. (Table 2; Fig 2)

Blood flow and pressures for the joint (ga^r, Pacir and P^cît) were 37.1 ml/min,

133.6 mmHg and 10.5 mmHg, respectively after joint isolation and during isogravimetric conditions. (Table 2) Capillary pressure was 22.6 mmHg and total vascular resistance was

13.5 mmHg/ml. The precapillary contribution to the total resistance was 73% (9.55 rmnHg) and the postcapillary (venous) contribution was 5.42% (0.71 mmHg). (Table 3) At isogravimetric conditions, Pcap significantly correlated with the isogravimetric venous pressure (Pv^r) (p<0.05). (Fig 3)

206 Lymph flow was zero and trans-synovial fluid flow was 5.28 drops/min or approximately 300ul/min during the isogravimetric state for the joint. (Table 3) The osmotic reflection coefficient was based on synovial fluid and was 0.22 for total protein and 0.18 for albumin. Lymph was only obtained from one horse and did not increase with

Pvcir. Lymph flow was not obtained due to immobilization of the joint preparation, maintenance of an lAP of zero and preferential trans-synovial fluid flow. The oncotic pressure exerted by synovial fluid (7.82 mmHg) was significantly greater than lymph (2.82 mmHg; p= 0.02) and less than plasma (13.3 mmHg). This resulted in a lower TMP for synovial fluid flow (mean 5.48 mmHg) than for lymph flow (10.7 mmHg; n=l).

Isolated Joint Vascular and Trans-synovial Forces and Characteristic Response to Pressure

Manipulation

Isolated joint blood flow (Qadr) significantly increased with Pa^r (range 100-250 mmHg; p= 0.002); peak 46.83 ml/min at Pacir 250 mmHg) and did not change with increased Pv^r. (P= 0.67; Table 2; Fig 4)

Measured joint vascular pressures (Pa^r and Pvcir) closely approximated the predetermined set value for the pressure manipulations (i.e., arterial pressures of 100,150,

200,250 and venous pressures of 20 and 40). (Table 2) Venous pressure of the joint at the isogravimetric state (PVcir= 10.5 nun Hg) was significantly lower than venous pressure of the entire distal limb (pre-preparation baseline; Pvcir = 20.0 mmHg). Pvcir did not significantly change as Pacir was increased (range 7.0 - 7.8 nunHg; p=0.38). Padr did not significantly change as Pvcir was increased (range 164 - 167.2 mmHg; p=0.62). (Table 2)

Joint vascular resistance (Pt) did not significantly change during arterial or venous pressure manipulations (p= 0.22)

207 Trans-synovial fluid flow (Qs) did not significantly increase with increasing Padr up to 200 mmHg (mean 4.08 drops/min [approximately 250 |il/min]; Table 2; Fig 5). At

Padr of 250 mmHg, trans-synovial fluid flow significantly increased (28.6 drops/min; approximately 1.73 ml/min) and returned to the previous flow when the joint preparation was returned to isogravimetric conditions (5.28 drops/min) (p=0.047). Synovial fluid production did not significantly increase with increased Pvdr (p= 0.24).

During the arterial pressure manipulations from isogravimetric conditions, the weight of the isolated joint significantly changed [as measured after the vascular phase of the weight gain curve] (p=0.028). (Fig 6) Weight significantly increased from Padr 100 mmHg to Padr 250 mmHg. (p=0.03). Weight did not significantly increase with increased

Pvdr. (Fig 6) Tissue compliance significantly increased as arterial pressure increased.

(P=0.05; Table 2; Fig 7)

Vascular compliance was low (overall mean 0.005) in the isolated joint during arterial pressure manipulations, but changed in direct concordance with changes in Padr-

As Padr increased from 100-250 mmHg, vascular compliance increased from 0.003 to

0.01. (P = 0.01; Table 2; Fig 8) The vascular compliance returned to 0.002 when the joint preparation was returned to isogravimetric conditions (121 mmHg). (Table 2; Fig 8) A marked increase in vascular compliance occurred from 0.002 to 0.08 during venous occlusion and increased Pvdr, and remained increased at Pvdr40 (p=0.0003).

Discussion

Our study is the first to describe a pump-perfused joint model that isolates the articular structures for physiologic investigation. This model successfully maintained physiologic function and synovial tissue perfusion for 4.5 hours. The quantity of tissue and fluid production was adequate for multiple sampling. Expected responses to

208 physiologie pressure manipulations occurred and added to our existing knowledge on the interrelation of vascular and codependent trans-synovial fluid shifts. For example, filtration pressure for synovial fluid production was mainly influenced by capillary pressure because of the low osmotic reflection coefficient and relatively high permeablity to proteins compared to other tissue beds.

The mild changes in systemic hemodynamics observed during the extended periods of general anesthesia were predictable.8.i7,5i.55The time-related changes in CO, PB.O2 and

PaC0 2 has been observed in horses under long term general anesthetic agents i? and may be related to alterations in autonomic tone ( CO) and positional related responses in animals of large body size.8.5i Controlled ventilation depresses cardiac output by decreasing venous return to the heart.25 Importantly, for anesthetized horses, all values were well above our minimum criteria and within the reported acceptable ranges; CO 30-60 L/min/kg,

Pa02 100-350 mmHg, and PaCOg 40-60 mmHg.24

Synovium blood flow of the MCP joints (0.073 ml/min/g) was similar to that reported for normal equine carpal joint synovium (0.1 ml/min/g) using a similar technique and did not significantly change throughout the duration of our study .22 Total blood flow to the equine digit has been similarly reported at 0.079 ml/min/g.2 Evidence of normal tissue perfusion was required to document minimal influence of the manipulations to create the preparation, and the duration of anesthesia. Normal tissue blood flow suggests minimal disturbances in vascular tone, vascular shunting and/or microvascular thrombus formation.

Our total limb flow was 68 ml/min and included flow to the digit and the MCP joint. The isolated joint demonstrated both hemodynamic similarities and differences to other isolated musculoskeletal organs, such as the digit and complete limb preparations.2.io.i3,46 The MCP joint blood flow was 37.1 ml/min at the Pacir of 133

209 mmHg during the isogravimetric state in lateral recumbency. When our joint arterial pressure was reduced to 100 mmHg, MCP joint blood flow decreased to 11 ml/min. Based on these findings, MCP joint blood flow was 54% of distal limb flow at an arterial blood pressure of 133 mmHg and decreased to 16% of total limb flow at an arterial blood pressure of 100 nunHg. An increase lAP would be expected to further decrease MCP joint blood flow, at least synovial flow, but this was not measured in our study .22 In a study of the isolated equine digit in a standing vertical position, total digit flow was 7.9 ml/min/lOOg and Pacir was controlled at 100 mmHg.2 The total digit preparation weights were not reported in the study.2 A comparison of our physiologic MCP joint values to those obtained for the isolated equine digit are listed in Table 3.2Our measured isogravimetric joint circuit pressure ( Pacir=133 mmHg) was within the physiologic range (80-150 nunHg)23 and was slightly less than the total limb pressure (149 nunHg). Other reports have measured equine distal digit pressures at ~ 100 nunHg.2 Based on these reports, pressure manipulations were selected from between 100 nunHg to 250 nunHg in order to assess changes within and beyond the physiologic range. Our measured isogravimetric venous pressure (Pvcir = 10.5 mmHg) and Pcap (22.6 nunHg) for the MCP joint was less than the Pv^r (30 mmHg) of the vertical digit.2The most obvious explanation for this difference is that our horses and our joint preparation were positioned in lateral recumbency. The high Pcap in the digit is countered by a high tissue pressure (Pt) and is recognized as being higher than analogous tissue pressures in dogs and humans.2.io.i3,i5,3i Our isogravimetric pressures identified for the joint approximate values in the canine limb.io.i3

Osmotic reflection coefficient using lymph could not be regularly measured in our study because lymph flow was absent at normal venous pressures. This was predictable since the limb was in lateral recumbency (no increase in tissue pressure as during

210 weightbearing). Lymph flow does occur from the equine limb, although in small quanity, when the animal is weightbearing and moving.5 Other studies have confirmed that lymph flow from normal joints is minimal.34.47 Lymph was obtained in one case but could not be accurately used to calculate Oj as it was not obtained at filtration independent lymph flow rates. Higher values for Oj hve been reported based on lymph values in other species’ joints37 and the equine digit (0.661 and -0.76). Our estimate of the osmotic reflection coefficient, an index of tissue permeability^ for the synovium (0.22), was much lower than for lymph (0.6 - 0.7) and reflected the higher total protein concentration in synovial fluid

(range 2.6-4.7 g/dl) at high flows. The synovial cavity is the dominant interstitial space in the joint and Od based on synovial fluid production more accurately reflects trans-synovial fluid forces. Other studies have reported higher Od (0.7-0.91) for tissues such as skin, intestine, and lung.56.59

Our findings, including a low Od partially explain why trans-capillary filtration produced synovial flow (trans-synovial filtration), rather than lymph flow. During isogravimetric conditions, the hydrostatic pressure gradient producing flow across capillaries to the synovial fluid (TMPs) was significantly less than for lymph (TMPi). This suggests a relatively high hydraulic conductance (i.e. the change in flow/unit change in driving pressure/area)38 for synovial tissue. The greater permeability of synovium (Od), and subsequently the greater concentration of protein in synovial fluid, facilitates osmotically driven fluid flux and filtration to the joint cavity in preference to lymph and has been suggested by other authors .35 Our documentation of this combination of physiologic data (higher synovial osmotic pressure, lower C7d and lower hydrostatic pressure threshold

211 for synovial fluid flow [TMPj]) explains our findings of active trans-synovial filtration without lymph flow, and threshold pattern of synovial fluid filtration with increased arterial pressure.34.39 a “yield phenomenon” has been described in studies of trans-synovial fluid absorption and occurred at intra-articular pressures of - 9cm H 2O (-7.3 mmHg).34The presence of a “yield or breaking pressure”3439 for filtration was apparent in our data since flow increased only after arterial pressure was 200 mmHg. The exact threshold pressure for inititating lymph flow was not investigated. Morphologic changes, including increased surface area and reduced capillary depth partially explain this effect for absorption of synovial flu id .3 9 Trans-synovial filtration (6$; synovial fluid production) is dependent on the hydraulic conductance of the membrane and the surface area of the o rg a n .5 6 In our study, the increased Qs with increased Pa was probably due to greater tissue hydraulic conductance, but possibly also increased villous capillary surface area during increased arterial pressures. Based on our data, it is probable that the TMP of synovium (TMPs =

5.48 mmHg) was exceeded at arterial pressure of 200 mmHg and drove fluid production.

Since lymph flow did not increase, the TMP of lymph was probably not exceeded.

Although synovial fluid production did not significantly increase with increased venous pressure, a trend was apparent (Fig 5), and probably represented a corresponding increase in capillary hydrostatic pressure. Increases in capillary pressure correlated closely with increased venous pressure in our study. (Fig 3) The increased capillary pressure probably exceeded the TMP for synovium when venous pressures of 20 - 40 mmHg were produced. Lymph flow did not increase at these venous pressures suggesting that the TMP for lymph was not exceeded. Increases in venous pressure in rabbit stifles produced a greater increase in outflow of injected oil than when arterial pressure was increased

(d<2s/?v > dQJP^, ratio 1.54).29 A similar ratio was not detected in our study, but greater

212 increases in venous pressure (> 40 mmHg) may have been required to produced larger increases in

Synovial flow should be high when determining Od so that protein concentration is not flow dependent. 3,18. Filtration independence was probably achieved in our study for synovial fluid since the high circuit pressures induced high synovial fluid flow rates. Our study documented that lymph flow is unlikely to achieve high flow states in joints with zero lAP when vascular pressure increases. Lymph flow may increase when lAP increased. Our studies suggest that an lAP of > II nunHg should have exceeded the

TMP\, and increased lymph flow. Levick reported that an lAP of 18 cm H 2O (-13.8 mmHg) minimally increased lymph flow .29.34 Pathologic joints with increased lAP however, demonstrate greater lymphatic clearance based on scintigraphy.47

Directional changes in MCT vascular forces in response to pressiue manipulations produced expected physiologic responses.29.33.46 Steady increases in joint blood flow produced the corresponding increased arterial pressure. (Fig 4) Increasing Pacir did not change Pvcir and increasing Pv^r did not change Pacir.29.33.46 (Table 2)

Our calculated total joint vascular resistance did not change significantly after each pressure manipulation, although the results were variable (Table 3). The precapillary component of vascular resistance composed the majority of the resistance (72%) similar to values reported for the equine digit (92 % ) .2 In our study, the calculated pre- and post capillary vascular resistance did not total 100% of the vascular resistance (Pt). The difference in pre- and post-capillary vascular resistance and the total vascular resistance of the joint probably reflected the resistance to fluid flow from the capillary to the synovial fluid (Ps) and accounted for 21.6% of Pt- Our study recorded that filtration exists from the capillary bed to the synovial fluid (-300 ul/min) when lAP is zero. Theoretically, if lAP

213 was increased enough to stop fluid flow to the joint cavity (based on our calculations of

TMPs, an lAP >11 mg Hg), then /?pre would increase its percent contribution to /?t as noted in digit preparations that do not have a steady fluid loss or lymph flow would increaseWe did not measure pre- and post capillary resistance at each pressure manipulation. As synovial fluid production increased with vascular pressure manipulations,

%Rs may have changed in proportion of Ri.

The immediate change in weight of the preparation after a pressure change suggests changes in vascular volume (ml/mmHg). The total length and area of the vascular bed is known to be proportional to the weight (in grams) of the isolated preparation. Furthermore, acute changes in weight (A W) caused by pressure changes are due to changes in area and length of the vessels.23r33.46 Associated changes in pressure (A P) and their relationship are represented by vessel compliance and can be estimated by A W/ A P. 2,23.42,50,52 This estimate of vascular compliance (AW/ AP) significantly increased as pressure in our studies increased suggesting that vessels become less stiff, (ie, more compliant). Vascular compliance (as estimated by weight changes in amputated perfused hindlimbs of rats) significantly increased with blood flow as in our study, however, an optimal flow was identified (0.25 ml/min) after which vascular compliance decreased.23 We did not identify an optimal flow for vascular compliance in the normal joint.

Our marked increase in vascular volume and compliance after increased venous pressure probably represents pooling of blood in venules. Arterioles are typically less compliant than venules, due to a myogenic response in the muscular layers of the arteriole wall.20,42 Because the method used to increase venous pressure also occluded (decreased) flow, accumulation of blood resulted in a large increase in vascular compliance. Studies using total limb preparations have demonstrated that increases in venous pressure by

2 1 4 venous occlusion produces the same change in limb weight as a 5- lOX greater increase in arterial pressure.46 Increase in vascular load by increasing blood flow would be expected to increase vascular volume and possibly alter vessel vascular function (compliance). Our study documented this to occur in the isolated joint to a small degree (Rg 8 ). Because venous flows correspondingly increased and no venous occlusion was present (ie, fv^r did not change), obstruction to outflow did not occur and vascular pooling of blood was minimal. As with any estimate of vascular compliance that uses weight change for the calculation, it is possible that vessel compliance did not change but that total vascular volume increased as a result of opening new vascular beds^z, although studies suggest changes in volume occurs initially.

Tissue compliance was estimated from the weight change associated with the hemodynamic pressure manipulations .20 Preparation tissue weight (g) was a sensitive indicator of fluid accumulation in our MCP preparation and was used to determine when the joint was in the isogravimetric condition (no loss or gain of fluid). The weight changes accurately reflected tissue weight changes because synovial fluid was removed and maintained at zero. Hydrostatic pressure in the c^illaries would be expected to increase with increased arterial pressure, and net fluid filtration would increase as documented in our study by an increase in synovial fluid flow. Fluid accumulated in the synovial tissue when MCP joint arterial blood pressures were between 200 and 250 mmHg and was detected as preparation weight gain. As tissue pressure increased and exceeded TMPs, synovial fluid production from the joint also increased. The synovial tissue became more compliant (accepted more fluid for a given pressure change) as pressure increased. (Fig 7)

Tissue compliance returned to baseline after the arterial pressures retumed to normal (120 mmHg) and an isogravimetric condition was established, although joint weight did not completely return to baseline. (Fig 6) Adequate time (30 minutes) may not have been

215 provided for complete clearance of the accumulated tissue fluid and may explain why a significant increase in preparation weight was not seen with further increased venous pressures as expected. (Fig 6)

In summary, our study suggests that the isogravimetric MCP joint has no lymph flow and a net filtration of synovial fluid of ~ 300 ul/min at zero lAP. Resistance to synovial fluid flow is responsible for approximately 21% of the total vascular resistance.

Arterial pressures > 200 mmHg are required to exceed the threshold arterial pressure for the joint and substantially increase synovial fluid flow. Trans-synovial flow occurs in preference to lymph flow due to the high hydraulic conductance and permeability of synovial tissue (low Od). Transitional microvascular pressures in excess of - 11 mmHg are needed to produce an increase in synovial fluid flow. Vascular compliance changes caused by increases in arterial pressure are minimal compare to those produced by increased venous pressure due to the greater elastance of arteries and the larger muscular arterial wall.

Synovial tissue becomes significantly more compliant as tissue fluid accumulates with the various vascular pressure manipulations and supports the conclusion that hydraulic conductance is high in synovial tissue. In conclusion, this isolated joint preparation permitted the evaluation of codependent hemodynamic, microvascular, and trans-synovial flow responses to hemodynamic manipulations.

216 Footnotes a-Masterflex variable speed pup. Cole Parmer Int, Chicago, IL b-Cole Parmer Int, Chicago, IL

C-FT03, Grass Instruments, Quincy, MA d-Grass Model 70 Polygraph, Grass instrument Co, Quincy, MA e-Spectramed P23XL transducers Oxnard, CA f-VR-12, Honeywell Corp, Pleasantville, NY g-Dye-Track, Triton Technology Inc, San Diego, CA h-Bio-rad Laboratories Inc, Richmond, CA

217 Figure A.l Pump-perfused isolated metacarpophalangeal joint preparation.

218 Pump Arterial pressure To VRI2

FT03 weight IL______a To physiograph Artenal circuit

Reference sample

Venous pressure

Median artery To physiograph Median vein Synovial fluid

Venous collection Venous circuit

219 Figure A.2 Equine forelimb and metacarpophalangeal joint before joint isolation demonstrating vascular sites for microsphere studies.

220 64 cm

Carpus Median artery Antebrachiocarpal joint Midcarpal joint

Medial palmar artery

Forearm g — Metacarpo phalangeal joint Reference syringe

221 Figure A 3 Capillary pressure versus venous pressure during isogravimetric conditions.

Capillary pressure (Pcap) was greater (p<0.05) at higher circuit venous pressure (Pvcir) [y

= I.4ÔX + 10.8; r = 0.87].

222 60 -,

50 -

b 3 i a 3 a 5

10 -

0 10 20 30 Cirait venous pressure (mmHg)

223 Figure A 4 Change in circuit arterial blood flow caused by manipulation of circuit arterial (Pacir) and venous (Pvdr) pressure (p < 0.(X)9). The first bar from the left (a 149) is before joint isolation, the second bar is after joint isolation at isogravimetric conditions.

Different letter superscripts are significantly different (p<0.05).

224 Circuit Blood Flow (ml/min)

Preprep

Postprep

Pal 00

0 Ë c TV -0 1 Pa200 g Pa250

Pal 20

Pv40 Figure A.5 Synovial fluid production (drops/min) caused by manipulation of circuit arterial (Padr) and venous (Pv^r) pressures (p<0.021). Different letter superscripts are significantly different (p<0.05).

226 a100 a150 a200 a250 a120 v20 v40 Arterial (a) or Venous (v) Circuit Pressures (Pair mm Hg)

227 Figure A.6 Preparation weight gain (g) caused by manipulation of circuit arterial (Pa^r) and venous (Pv^r) pressures (p< 0.04 Pa^r; p < 0.57 Pvcir). Different letter superscripts are significantly different (p<0.05). Letters with a “t” superscript tended to be different ( p<0.1).

228 (D (.03) p:.02)

a 100 also a 200 a 250 a 120 v20 v40 Circuit Pressures (mm Hg)

229 Figure A.7 Tissue compliance (ml/mmHg) changes caused by arterial (Padr) (p<0.05) and venous Pv^r pressure manipulation. Different letter superscripts are significantly different (p<0.05).

230 0 .0 9 -

0.08“

E 0 .0 7 -

E 0 .0 6 - 8 ^0.05- Q. I 0 .0 4 - Ü ® 0 .0 3 -

0 .0 2 -

0 .0 1 -

a150 a 200 a 250 a 120 v20 v40 Arterial (a) or Venous (v) Circuit Pressures (P cir mmHg)

231 Figure A.8 Vascular compliance (ml/mmHg) changes caused by arterial (Pacir) and venous (Pv^r) pressure manipulation (p<0.01). Different letter superscripts are significantly different (pcO.05). ^ = a trend ( p<0.1) to be different from Pv20. Letters marked with a “t” superscript tended to be different (p<0.1).

232 0 .0 9 -

0 .0 8 -

0 .0 7 -

0 .0 6 -

0 .0 5 -

0.04-

0.03-

0.02-

0.0 1 -

1------1------1 ' I------1 ‘ I a 150 a 200 a 250 a 120 v20 v40 Arterial (a) or Venous (v) Circuit Pressures (Pcir mmHg)

233 RRRF.D AGE (vrs) SEX BODY WEIGHT!k-.

QH ty p e adult MC 400

QH 1.5 MC 306 TB 1.0 MC 320

QH 2.0 F 402 TB 4.0 MC 427 TB 1.0 M 355 TB 5.0 F 556

MC= male castrated; M= male; F= female; TB= Thoroughbred; QH= Quarter Horse

Table A .l Signalment of horses used in the isolated metacarpophalangeal joint preparation (n=7).

234 1 Pfo oroo basetine Poat'Or#D basalln# ' P a 100 ImmHo) 1 Pa 150 (mmHa) Pa 200 (mmHo) P a 250 (mmHo) Pa 120 ImmHo) Pv 2 0 (mmHo) Pv 40 (mmHo)

Cardiac output (L/min/ttg) 0 07a +A 0 011 0 006 * h 0.009 0 000 W* 0 011 0 050 «A 0 004 0 055 4/' 0 005 0 050 tA 0 004 0 05 tA 0 003 0 049 tAO 004 0 0451 «/ 0 004 w#an Aft#mat P rtstur# (mmHg) 130.07 +A 18 23 123 80 ♦/• 110 2 136.76 140 0 105 6 «A20 06 12707 4/. 11 2 124 0 tA 11 7 132 tA 9 30 121 33 tA 1028 1)2 5 ,r 10 27 Sydoitc Artertal Prttaur# 100.03 */ 22.17 1820 tA 14 24 104 00 4A 17 40 100 67 «/• 1251 157 17 4/. 12 4 151 5 tA 10 0 101 0 tA 14 22 140 17 t / 14 32 1420 */ 13 35 Aftonal Btood Oa# (Pa02) 422.00 *!• 30 ao 401 03 *A 36 3 395 8 0 439 7 tAO 359 6 tA 0 (PaC02) 40.48 2 00 37 4 */. 3.38 43 0 40 1 tAO 54 8 tAO Joint Blood FLom (mi/min) 08 21 13 37.17 13 05 11.0 +A 4 0 8 14 8 »/• 5 23 24 07 4A 7 05 40 83 tA 10 95 10 33 tA 541 20 42 tA 7 50 20 42 tf 603 Joint ArtanaJ Prtaaura (mmHg) 149 47 12 74 13357 W- 9 07 108 +A503 154 83 */• 2 57 201 07 tA 2201 249 0 t/. 1 121 6 tA 9 49 164 tA 1886 167 17 1/ 1650 Jomi Vaftoua P rttaur# (mmHg) 20 0 W'O 10 5 W. 3 81 7 33 W. 1 33 7 0 */• 1 33 7 83 4A 2 20 7 0 tA 1 80 9 0 tA 2 90 20 0 tA 0 00 40 0 */ 0 00 Synowwm Blood Flow (mUmirVg) ■doraaJ 0.078 W- 0 047 0.058 +/. 0 009 0 049 tA 0 03 •palmar 0 007 W- 0 034 0.201 4/. 0.118 0 048 ♦/• 0 014 •moan 0.073 +/. 0.000 0 160 ♦/« 0 102 0 049 tA 0 001 Jomt Raaiftance (Rl) mmHgfml 2.27 ♦/. 0 61 0 41 *A 2 02 12.01 ♦/• 6 07 17 78 ♦/. 6 4 11 53 4A 2.74 10 89 tA 3 54 13 09 tA 4 27 17 34 tA 6 66 13 35 ,/ 531 Prap weight gain (g) 4.33 4/. 3 86 4 0 »A 2 58 10 0 4/. 7 32 23 07 tA 840 18.0 tA 9 17 168 tA 4 54 12 83 tA 5 65 Synovial Fluid Production (dropaAmn) 1 07 4/ 0 07 4 08 */• 1.42 4 08 4A 2.37 28 57 tA 13 0 5 28 tA 2 44 8 73 tA 4 74 14 74 tA 1045 Vascular CompHane# (mVmmHg) 0.042 4/. 0 002 0 004 */. 0 001 0.008 4A 0 002 0 010 4/' 0 003 0.003 W* 0 001 0 061 tA 0 033 0 039 t/ 0013 Tissue comoliance fmMnmHo) 0 0114 «/• 0 00 0 053 4A 0 011 0 074 tA 0 010 0 025 4/ 0 007 0 02 tA 0 004 0 020 «A 0 010 s

Table A.2 Systemic hemodynamics and isolated joint vascular function (mean ± SEM) in 7 horses. Reported values for Measured Variables Values Obtained equine digit

Joint Arterial Pressure (Pacir mmHg) 121.5 +/- 9.49 100.25 +/- 5.12 Joint Venous Pressure (Pvcir mmHg) 9.0 +/- 2.90 27.37 +/- 4,85 Joint Capillary Pressure (Pcap mmHg) 22.60 +/- 4.07 36.67 +/- 1.79 Joint Resistance (Rt mmHg/ml) 13.09 +/- 4.27 8.00 +/- 1.05 Joint Arterial Resistance (Rpre mmHg/ml) 9.55 +/- 3.66 7.43 +/- 1.03

Joint Venous Resistance (R post mmHg/ml) 0.71 ~ l - 0.38 0.657 +/- 0.066 Pre- to postresistance ratio 13.45 12.41 +/- 2.68 Contribution R pre (%) 72.96 9 2 .8 8 Contribution R post (%) 5.42 8.21 Osmotic Reflection Coefficient -Total Protein 0.22 +/- 0 -Albumin 0.18 +/- 0 . Oncotic Pressure Synovial Fluid (its) mmHg 7.82 +/- 2.9 Oncotic Pressure Plasma (up ) mmHg 13.31 +/- 3.7 19.09 +/- 0.84 Oncotic Pressure Tissue ( itt) mmHg 2.82 +/- 0 6.59 +/- 1.54 Net nitration Pressure [Pcap+its-itp] 17.1 +/- 4.05 Net Filtration Pressure [(Pcap)-0d(itp-its)] 21.4 +/- 4.05 Vascular Compliance (ml/mmHg) 0.003 +/- 0.001 0.012 +/- 0.002 Tissue Compliance (ml/mmHg) 0.025 +/- 0.007

Table A J Isolated metacarpophalangeal joint vascular and fluid values (mean ± SEM) at isogravimetric conditions.

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