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The Journal of Immunology, 2001, 166: 370-376.
Copyright © 2001 by The American Association of Immunologists

T Lymphocytes Promote the Development of Bone Marrow-Derived APC in the Central Nervous System1

Sandhya Subramanian*, Dennis N. Bourdette*,{dagger}, Christopher Corless*,{ddagger}, Arthur A. Vandenbark*,{dagger}, Halina Offner{dagger} and Richard E. Jones2,{dagger},{ddagger}

* Veterans’ Affairs Medical Center, Portland, OR, 97201; {dagger} Department of Neurology, Oregon Health Sciences University, Portland, OR 97201; and {ddagger} Oregon Cancer Center, Portland, OR 97201


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Certain cells within the CNS, microglial cells and perivascular macrophages, develop from hemopoietic myelomonocytic lineage progenitors in the bone marrow (BM). Such BM-derived cells function as CNS APC during the development of T cell-mediated paralytic inflammation in diseases such as experimental autoimmune encephalomyelitis and multiple sclerosis. We used a novel, interspecies, rat-into-mouse T cell and/or BM cell-transfer method to examine the development and function of BM-derived APC in the CNS. Activated rat T cells, specific for either myelin or nonmyelin Ag, entered the SCID mouse CNS within 3–5 days of cell transfer and caused an accelerated recruitment of BM-derived APC into the CNS. Rat APC in the mouse CNS developed from transferred rat BM within an 8-day period and were entirely sufficient for induction of CNS inflammation and paralysis mediated by myelin-specific rat T cells. The results demonstrate that T cells modulate the development of BM-derived CNS APC in an Ag-independent fashion. This previously unrecognized regulatory pathway, governing the presence of functional APC in the CNS, may be relevant to pathogenesis in experimental autoimmune encephalomyelitis, multiple sclerosis, and/or other CNS diseases involving myelomonocytic lineage cells.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Multiple sclerosis (MS)3 is a debilitating paralytic inflammatory disease of the CNS. Experimental autoimmune encephalomyelitis (EAE) is a paralytic inflammatory disease induced by myelin-specific T cells in rodents (1, 2, 3). Although the cause of MS is not known, clinical and pathologic similarities between MS and EAE suggest that MS may be an autoimmune disease mediated by myelin-specific T lymphocytes in the CNS (4, 5, 6, 7).

The pathogenicity of myelin-specific T lymphocytes has been confirmed in EAE by adoptive transfer of characterized T cells (8, 9, 10, 11, 12, 13, 14, 15, 16). In order for circulating myelin-reactive T cells to induce CNS inflammation they must enter the CNS tissue parenchyma (10). In the CNS, a barrier between the blood and the CNS tissue parenchyma, the blood-brain barrier (BBB), is formed by vascular endothelial cells, the basement membrane, CNS microglial cells, and astrocytes (17, 18). Circulating T cells expressing an activated phenotype are able to cross the BBB and enter the CNS (19), whereas circulating naive T lymphocytes expressing a resting phenotype cannot readily cross the BBB (17). T cell entry into the CNS does not depend on the T cell’s specificity for neural Ag. However, activated T cells specific for myelin Ags persist for a longer time and induce inflammation within the CNS (20). Thus, the activation state of circulating T cells controls their entry into the CNS, and Ag specificity (for myelin Ag) determines whether T cells will be stimulated to express a pathogenic phenotype in the CNS.

Following their entry into the CNS, further stimulation of Ag-specific CD4+ T cells depends on a specific molecular interaction between the TCR and the complex of syngeneic MHC class II and peptide Ag (21) expressed on the surface of specialized APCs (22, 23, 24). For this reason, encephalitogenic, myelin-specific T cells are unable to induce EAE following T cell transfer into irradiated allogeneic recipients where syngeneic APC are not present (25, 26). However, syngeneic APC derived from transplanted bone marrow (BM) could be introduced into the CNS of allogeneic recipient rats, appearing in brain and spinal cord (SC) tissue within 2–6 mo following BM transplantation (25, 26, 27). Subsequently, it was possible to transfer EAE into these chimeric allogeneic recipients, provided that the T cells were syngeneic with the transplanted BM and were specific for myelin determinants expressed in the CNS (25, 26).

In vitro similarities between encephalitogenic rodent T cells and certain myelin-reactive T cells from MS patients (13, 28, 29, 30) indicate that at least some human T cells are probably pathogenic in MS. However, a method has not yet been developed to distinguish whether or which human T cells might be encephalitogenic. The ability to perform adoptive transfer experiments with characterized human T cells in chimeric mice would facilitate the identification and characterization of the distinct T cells that cause MS and could provide an in vivo model for testing treatments directed against disease-causing human cells (8, 9, 10, 12, 31, 32, 33). In principle, BM transplantation should allow the construction of chimeric mice that are susceptible to EAE mediated by a variety of histoincompatible T cells (25, 26), including, as we have previously proposed (34), those from human donors.

The in-bred SCID mouse strain C.B-17scid/scid is deficient in both T and B lymphocytes (35) and lacks the immune responses associated with these cells. SCID mice have been shown to be suitable recipients of functional human BM and blood cells (36, 37, 38, 39). Therefore, for the current report we have chosen to use SCID mice as BM and T cell recipients to examine the requirements for inducing EAE in chimeric mice with nonmurine T cells, anticipating that these methods might be adaptable to the use of human T cells in the future. These experiments were conducted using a known EAE-susceptible donor rodent strain (Lewis rat) as the source of nonmurine BM and encephalitogenic T cells.

Transplanted hemopoietic-derived Lewis rat cells populated the SCID mouse CNS with functional APC 4–6 wk after transplantation (40). Coincident with the development of rat APC in the CNS, SCID mouse-rat hemopoietic cell chimeras were susceptible to EAE induced by myelin-specific rat T cells (40). Thus, as with allogeneic BM transplant studies conducted previously in rats (25, 26), transplanted rat BM-derived APC populated the SCID mouse CNS very slowly, over a period of months, and functioned in the induction of EAE. To evaluate possible interactions between transferred T cells and developing APC, xenogeneic rat BM and rat T cells were transferred alone or together into SCID mice, and the resulting chimeric mice were evaluated for susceptibility to EAE and for the appearance, phenotype, and function of BM-derived rat cells in the CNS.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals

Six- to 8-wk-old female Lewis rats were obtained from Harlan Sprague-Dawley (Indianopolis, IN). In-bred female SCID mice (8–11 wk old) were obtained from the SCID mouse breeding facility within the Veterinary Medical Unit at the Veterans’ Affairs Medical Center. Animal care was in accordance with institutional guidelines.

Rat T cell lines

Eight rats per line were each immunized by s.c. injection with a synthetic peptide of myelin basic protein (BP) corresponding to amino acid residues 87–99 (BP87–99), chicken OVA (100 µg Ag/rat), or no Ag in CFA containing 100 µg heat-inactivated Mycobacterium tuberculosis strain H37RA (Difco, Detroit, MI). Nine days following immunization, draining lymph nodes (LN) were collected, and a single-cell suspension of LN cells was prepared. LN cells were stimulated in vitro (8 x 106 cells/ml) for 3 days with Ag (BP87–99, chicken OVA, or the purified protein derivative of M. tuberculosis (PPD; 200 µg/ml), depending on the desired specificity of the line. Stimulated cells were expanded in rIL-2-containing medium for 5–7 days. This was followed by restimulation of the T cells with Ag in the presence of irradiated Lewis rat thymocytes for 3 days. Cells were selected and maintained through alternate cycles of stimulation with Ag and expansion in IL-2. Ag specificity was verified for each cell line during Ag stimulation using an in vitro thymidine uptake proliferation assay, and the disease-inducing potential of the T cell lines was confirmed by adoptive transfer of activated T cells (10–20 x 106 cells per rat, i.p. injection) into Lewis rats (data not shown).

Cell transfer into SCID mice

The indicated number of Lewis rat T cell line cells were injected (0.2 ml/mouse) by i.v. injection into anesthetized irradiated (300–350 Gy) SCID mice. T cells were injected either alone or within 5 min of Lewis rat, T cell-depleted BM cells (BMC). BMC were injected i.v. (0.2 ml/mouse) either alone or within 5 min of the injected T cells.

Isolation of Lewis rat BMC

All experiments involving rat BMC used T cell-depleted BMC. Two or three adult female Lewis rats were used as BM donors. The femurs and tibias were dissected free of surrounding tissue and collected in ice-cold RPMI 1640 medium. After cutting off the ends of each bone, BMC were flushed out with ice-cold RPMI 1640 using a 1-ml syringe. BMC were pelleted by centrifugation, resuspended in 10 ml ice-cold RPMI 1640, counted, and then incubated with a 1:160 dilution (final) of mouse anti-rat CD3 mAb (PharMingen, San Diego, CA) for 30 min on ice. Following this, cells were washed and incubated with goat anti-rat IgG Ab-coated magnetic beads (Miltenyi Biotech, Auburn, CA). Cells were then washed and run through a magnetized column (Miltenyi Biotech) that had been pretreated with PBS/4% BSA for 30 min. The column was washed five to six times with staining medium (PBS/3% bovine serum) to elute all unbound CD3-negative cells. The column was then demagnetized by removal from the magnet, and the CD3-positive cells were eluted from the column. The total number of BMC recovered (CD3-negative plus CD3-positive BMC) from the magnetic column depletion procedure was 80–95% of the BMC that had passed into the column. Undepleted, T cell-depleted, and T cell-enriched BM fractions were stained with mAbs specific for rat T cells to verify the purity of the respective populations and to evaluate the effectiveness of the magnetic sort. After T cell depletion, it was not possible to detect T cells in the T cell-depleted BMC fraction by fluorescent mAb staining and flow cytometry using Abs specific for rat TCR, CD3, or CD5 (data not shown). T cell-depleted rat BMC were injected i.v. (0.2 ml) into anesthetized SCID mice.

Evaluation of clinical disease severity (paralysis)

Mice were evaluated daily for disease and scored as follows: 0, normal; 1, limp tail or mild hind limb weakness; 2, moderate hind limb weakness or mild ataxia; 3, moderately severe hind limb weakness; 4, severe hind limb weakness or moderate ataxia; 5, paraplegia with no more than moderate forelimb weakness; 6, paraplegia with severe forelimb weakness or severe ataxia.

SC cellular phenotyping

SC cells from SCID mice were evaluated for the presence of rat cells using fluorescent mAbs (PharMingen) specific for rat myelomonocytic cells (CD11b/c), T cells (CD3), and rat MHC class II (RT-1B). A single-cell suspension was prepared from freshly isolated SC. The spinal column was dissected from euthanized donor mice. The SC was expelled from the column with ~10 ml cold saline introduced under pressure from a 10-ml syringe with an 18-gauge needle inserted into the caudal end. The whole SC were pressed individually through a fine wire-mesh screen, and a single-cell suspension was collected and pelleted by centrifugation. The pelleted cells from each SC were resuspended in 5 ml 80% Percoll (Pharmacia, Piscataway, NJ) and overlaid with 5 ml 40% Percoll in a 15-ml centrifuge tube. Cells were centrifuged for 30 min at 1600 rpm, and cells from the 80/40 interface were collected. Cells were washed, counted, and aliquoted for mAb staining. Cells were incubated with fluorescent mAbs (1 x 105 cells/0.2 ml/tube) for 20 min on ice. Reactivity with specific mAbs was evaluated by flow cytometry using a FACScan (Becton Dickinson, Mountain View, CA). Dot plots of fluorescence intensity for FITC- and PE-labeled cells were evaluated using CellQuest software (Becton Dickinson) to identify and quantitate distinct populations of cells. Isotype-matched (control) fluorescent Abs were used at the same concentration as each specific Ab for every experiment. Percentage of true positive staining was determined by subtracting the percentage of the isotype control staining (background) from the percentage of staining of each specific mAb.

APC function of SC cells

SC cells from SCID mice were evaluated for their ability to stimulate Ag-specific rat T cells in vitro. A single-cell suspension of SC cells was isolated and pooled from five euthanized donor mice 8 days after injection of Lewis rat BMC and OVA-specific Lewis rat T cells using methods essentially identical with those described above for the isolation of CNS cells for phenotyping. SC cells or Lewis rat thymocytes were washed, counted, and resuspended at 2.5 x 105 cells/ml stimulation medium (RPMI 1640 (Life Technologies) containing 2% (v/v) autologous rat serum, L-glutamine, and 2-ME). Cells were added (100 µl/well) to 96-well, round-bottom microtiter plates. To each well, BP-specific T cells were also added (2 x 103 T cells/100 µl/well). Ags were added (no Ag (control) or 10 µl/well of a 1-mg/ml solution of BP87–99 or OVA), and plates were incubated for 72 h, 37°C, 7% CO2. Cells were labeled with 0.5 µCi [3H]thymidine/10 µl/well for the last 18 h of culture. Wells were harvested, and cpm were determined using a liquid scintillation counter (Betaplate 1205; Wallac, Gaithersburg, MD). Results are expressed as mean cpm ± SD as an indication of thymidine uptake and T cell proliferation.

Immunohistochemical staining of SC

Mouse mAbs specific for rat CD45 (IgG1), RT-1B (IgG1), and isotype-matched control Abs were purchased from PharMingen. The Abs (specific and isotype control) were diluted 1/500 in dilution buffer (10 mM PBS (pH 7.4/1%), BSA/0.05% Tween 20/0.1% sodium azide). Biotinylated anti-mouse IgG (Vector BA-2000; Vector Laboratories, Burlingame, CA) was used as the secondary Ab at a final dilution of 1:400.

Five-micrometer cryostat sections of the snap-frozen SC tissue were prepared on Capillary Gap Plus slides (Fisher Scientific, Pittsburgh, PA) and allowed to air-dry for 10 min at room temperature. The sections were wrapped in cellophane to protect them from condensation and stored at -70°C until used. Stored slides were warmed to room temperature, unwrapped, and immediately fixed in 1% paraformaldehyde in PBS for 10 min (room temperature). After two washes in PBS/0.05% Tween 20 (Sigma, St. Louis, MO), the slides were loaded onto a Biotek Techmate 1000 (capillary-gap type) automated immunostainer (Ventana Medical Systems, Tucson, AZ). After a 10-min incubation with dilution buffer, the slides were exposed to primary Ab for 45 min at room temperature. Following three washes in PBS/Tween 20, the slides were incubated for 30 min with anti-mouse IgG secondary in dilution buffer. After additional washes, the slides were incubated in a quench solution consisting of 80% methanol/3% hydrogen peroxide/1% sodium azide. The slides were again washed with PBS/Tween 20 and then incubated for 30 min with avidin-biotin complex (Vectastain Elite ABC kit; Vector Laboratories) prepared per kit instructions. After three final washes, staining was visualized by incubating for 10 min in diaminobenzidine solution (K3466; Dako, Carpinteria, CA) prepared per manufacturer’s instructions), after which the slides were rinsed in water, counterstained with Gill’s hematoxylin, dehydrated, and coverslipped.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
We transferred rat BMC alone or with rat T cells into SCID mice to determine the cellular requirements for development of EAE in chimeric SCID mice. The simultaneous cotransfer of 5–40 x 106 BP-specific Lewis rat T cells and 10–40 x 106 T-depleted, adult Lewis rat BMC into SCID mice induced an acute ascending paralysis (Table IGo). Eighty-one percent of the mice (51 of 63) developed clinical EAE (paralysis). Both BP-specific T cells and BMC were required for EAE because neither T cells alone nor BMC alone caused disease. Disease depended on T cell specificity for myelin because T cells specific for nonmyelin Ags (PPD or OVA) were not capable of causing disease following cotransfer with BMC (Table IGo).


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  		Table I. Clinical outcome following adoptive cotransfer of BP-, PPD- and OVA-specific rat T cells and/or T cell-depleted rat BMC in SCID mice1

 
Paralysis lasted 1–4 days and was followed by complete recovery (Fig. 1Go). The severity of disease was related to the number of cells transferred. Mice that received a total of 40 x 106 T cells plus 40 x 106 BMC (40/40, Fig. 1Go) developed a more severe disease than mice that received 10 x 106 T cells plus 10 x 106 BMC (10/10, Fig. 1Go and Table IGo). The onset of clinical EAE occurred as early as 8 days following cell transfer.



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  		FIGURE 1. Clinical course of paralysis in SCID mice. Multiple groups of SCID mice, from Table IGo, received 40 x 106 (40/40), 20 x 106 (20/20), or 10 x 106 (10/10) of each cell type, Lewis rat encephalitogenic T cells and T cell-depleted Lewis rat BMC. Mice were followed for 20 days and scored daily for the severity of neurologic deficit (paralysis, clinical EAE). Results are presented as the daily mean clinical disease scores (paralysis) of all animals that developed EAE (numerator of clinical disease incidence, Table IGo). Variations in the disease course (e.g., the day on which maximal disease occurred) between individual animals are reflected as differences between the maximal mean severity of disease achieved for each group in Fig. 1Go and the mean maximal severity of disease listed in Table IGo.

 
The kinetics of circulating BM-derived cells moving into the CNS was evaluated in the absence or presence of activated T cells specific for nonmyelin or myelin Ags (Fig. 2Go). Transplanted rat T cells entered the SCID mouse SC within 3–5 days following transfer of 40 x 106 OVA- or BP-specific T cells alone (Fig. 2Go, A and B, respectively). Unlike T cells, BM-derived, CD11b-positive rat cells were undetectable in the SC during the first 10 days following the transfer of 40 x 106 rat BMC alone (Fig. 2GoC).



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  		FIGURE 2. Rat cells in the SC following transfer of rat T cells and/or rat BMC. Two or three groups of SCID mice per experiment received by i.v. injection 40 x 106 of each indicated cell type (T cells and BMC, BMC alone, or T cells alone; day 0). Each experiment included one group of animals receiving both T cells and BM-derived cells. At the indicated time points, single animals from each group were examined concurrently for the phenotype of cells present within the SC. Results are expressed as cells per cord. This was determined for each cord at each time point by multiplying the percentage of cells positive for a particular cell phenotype marker by the total number of cells isolated per cord. The results shown are representative, incorporating data from multiple overlapping independent cell-transfer experiments, and have been verified in replicate experiments. "No EAE" signifies cell-transfer groups in which EAE was not induced. "+EAE" signifies the cell transfer group in which EAE was induced.

 
When rat T cells and BMC were transferred together, 2,000–10,000 T cells per cord were detected within 3–7 days after cell transfer, followed by the appearance of >3,000 BM-derived cells (CD11b-positive) per cord within 7 days (Fig. 2Go, D and E). T cells entered the CNS of individual mice earlier than BM-derived cells, and BM-derived cells entered the CNS earlier after co-transfer with T cells than after BMC transfer alone.

The first CD11b-positive cells to arrive in the CNS 7 days after the cotransfer of T cells and BMC did not express MHC class II (RT-1B) (Fig. 2Go, D and E, day 7). As early as 8 days following cell transfer, RT-1B was expressed on a substantial number (>2,000 cells per cord) of CD11b-positive cells (Fig. 2Go, D and E). Expression of RT-1B did not depend on Ag recognition by T cells, because limited expression occurred even in the absence of myelin-specific T cells (Fig. 2GoD). However, enhanced expression of RT-1B on the majority of CD11b-positive cells occurred within 10 days after co-transfer in the presence of myelin-specific T cells and was associated with disease (Fig. 2GoE). With nonmyelin-reactive T cells (specific for PPD), for which stimulating Ag was not available in the CNS, RT-1B expression on CD11b-positive cells remained relatively low on day 10 (2% of total cells {approx} 8% of CD11b-positive cells, Fig. 3GoA) compared with myelin-specific T cells (23% of total cells {approx}65% of CD11b-positive cells, Fig. 3GoB). RT-1B was also expressed on a minor population of CD11b-negative cells following transfer of activated PPD-specific (4% of total cells, Fig. 3GoA) or BP-specific (14% of total cells, Fig. 3GoB) T cells and BMC.



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  		FIGURE 3. Phenotype (A and B) and in vitro APC function (C) of SCID mouse SC after transfer of rat T cells and rat BMC. The phenotype of SC cells isolated 10 days after transfer of rat BMC and nonencephalitogenic, PPD-specific (A) or encephalitogenic, BP-specific (B) rat T cells is shown for representative SCID mice (compare to Fig. 2Go, D and E, respectively). BM-derived cells were detected with fluorescent mAbs specific for rat MHC class II (x axis) and rat CD11b (y axis). Fluorescence intensity of Ab-labeled cells is indicated on the dot plot. Some rat T cells (CD11b-negative) were positive for expression of rat MHC class II (4% and 14%). Values shown for percentage of positive cells labeled with specific Abs were calculated by subtracting the percentage of nonspecific labeling with isotype-matched, control fluorescent Abs (<4%). APC function of SC cells was tested in vitro (C). SC cells were isolated from SCID mice 8 days after transfer of Lewis rat OVA-specific T cells and Lewis rat BMC (mouse OVA SC). T cell stimulation responses were measured as the mean cpm (±SD) [3H]thymidine incorporated in triplicate wells during the final 18 h of a 72-h culture of BP-specific T cells (BP T) and APC without (control) or with Ag (BP87–99 or OVA). The response with rat thymus APC is shown for comparison.

 
Eight days after the co-transfer of rat OVA-specific T cells and rat BMC, cells were isolated from the SCID mouse SC and tested for their ability to present Ag to rat BP-specific T cells in vitro. BP87–99 peptide-specific T cells (BP T, Fig. 3GoC) proliferated in response to BP87–99, but not to OVA presented by pooled APC isolated from the SC of five chimeric SCID mice (mouse OVA SC, Fig. 3GoC). The specific T cell response was comparable to that using Lewis rat thymocytes as a positive control source of rat APC (rat thymus).

The co-transfer of myelin-specific rat T cells and rat BMC together caused a marked inflammation typical of EAE in the SC of SCID mice (Fig. 4Go). Compared with healthy naive SCID mice (Fig. 4Go, A and B), representative paraffin-embedded SC sections from SCID mice with EAE had mononuclear cellular infiltrates in the white matter 10 days following cell transfer (Fig. 4Go, C and D). The location and phenotype of rat cells in the SCID mouse CNS were evaluated using immunohistochemical staining of frozen SC sections to establish a tissue or vascular location and phenotype for the infiltrating rat cells. The rat-specific mAbs used did not react with frozen healthy mouse cord (Fig. 4GoE), and isotype-matched control Ab failed to react with diseased mouse cord (Fig. 4GoF). Eight days following the transfer of BP-specific T cells and BMC, the SC from a representative SCID mouse showed predominantly focal infiltrates of CD45+ (Fig. 4GoG) and RT-1B+ (Fig. 4GoH) rat cells, consistent with an intravascular and perivascular localization within the CNS tissue, similar to early EAE in non-chimeric mice. Two days later, 10 days following cell transfer, CD45+ (Fig. 4GoI) and RT-1B+ (Fig. 4GoJ) cells were more diffuse within the SC white matter of a paralyzed SCID mouse, with some foci of inflammatory cells remaining. Thus, during the induction of EAE in SCID mice, infiltrating rat leukocytes (CD45+) moved into an extravascular location (perivascular and diffuse) within the CNS tissue parenchyma and expressed RT-1B, the phenotype of APC and activated T cells.



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  		FIGURE 4. Inflammation and cellular infiltration in SCID mouse SC. Paraffin sections from the SC of a naive SCID mouse (A and B) and from a SCID mouse with EAE (C and D) are shown. Paraffin sections were stained with Luxol fast blue, periodic acid-Schiff’s reagent and hematoxylin. Inflammation in the white matter (stained blue) is visible as darkly stained nuclei at low and high power magnification (x50 and x200, respectively, before printing from a 35-mm slide). Frozen sections from SCID mouse SC were stained with hematoxylin and examined by immunohistochemical staining (E–J). A healthy naive SCID mouse SC section stained with a mAb (OX-1) specific for rat CD45 (E) failed to show reactivity, as expected for mouse SC in the absence of rat cells. Similarly, a SC section from a SCID mouse with EAE, 10 days following cell transfer (F), failed to react with isotype-matched (IgG1) "control" mAb at the same concentration as the OX-1 and MHC class II Abs. Eight days following transfer of rat BMC and rat encephalitogenic T cells, the SC of SCID mice reacted with the rat CD45 Ab (G) and the rat MHC class II Ab (H). Rat CD45 (I) and rat MHC class II (J) were also detected in the SC of SCID mice with EAE 10 days following cell transfer.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Based on prior studies in which transplanted rat BM-derived cells were shown to populate the CNS over a period of months in the absence of activated T cells (25, 26, 27, 40), the onset of clinical EAE within 8 days following the transfer of myelin-specific rat T cells and rat BMC in SCID mice was several weeks earlier than expected. Because encephalitogenic rat T cells require rat APC in the CNS to induce disease, this rapid onset of disease demonstrated that transplanted BMC require only 8 days to move into the CNS and develop into functional APC. Moreover, the results indicate that the selective barrier to migration of BMC across the BBB is altered by nonencephalitogenic (OVA- or PPD-specific) as well as encephalitogenic (BP-specific) activated T cells. Accelerated entry of BM-derived APC into the CNS, mediated by activated T cells in the absence of reactivity with myelin Ags, represents a previously unrecognized pathway governing susceptibility to CNS inflammation. Through such a pathway, it is possible that activated T cells involved in responses to infectious pathogens could potentiate an encephalitogenic event mediated by myelin-specific T cells. We are presently initiating studies to quantitate the disease-potentiating activity of nonmyelin-specific T cells in SCID mice.

Previous studies of EAE in rodent BM chimeras suggested that BM-derived cells residing in the CNS were necessary before the arrival of encephalitogenic T cells (25, 26, 27, 34, 40), and a substantial disease-initiating role for circulating BM-derived APC was not demonstrated. In the current study, susceptibility to EAE induced by rat T cells did not require resident, functional rat APC in the SCID mouse CNS before the transfer of encephalitogenic rat T cells. Rather, the transferred T cells mediated a rapid acute migration of circulating BM-derived cells to the CNS before the onset of disease, and these BM-derived cells were entirely sufficient to supply the APC functions necessary for causing inflammation and disease. Thus, in the presence of activated T cells, the role of circulating BM-derived APC in the initiation and development of CNS inflammation may be greater than previously thought. Given the crucial role for APC in CNS inflammation, it will be important to determine the relative contributions of CNS-resident APC and T cell-recruited APC.

The results also indicate that an elevated expression of MHC class II on the majority of CD11b-positive cells depended on the T cells’ pre-existing potential for activation by neural Ag plus APC and suggest that signals originating from neural Ag-activated T cells were responsible for the observed increased expression of MHC class II on CD11b-positive cells during the early stages of disease induction. However, the results do not distinguish between up-regulation of MHC class II and recruitment of new MHC class II-positive cells as possible mechanisms to explain increased levels of MHC class II expression by CD11b-positive cells in the CNS.

MHC class II was also up-regulated on CD11b-negative cells. These cells most likely represent rat T cells expressing an activated phenotype (41, 42) as a result of the in vitro stimulation with Ag before transfer into SCID mice or, in the case of BP-specific T cells, as a result of further activation due to in vivo stimulation by APC and Ag. The activated phenotype has been shown previously to be required for T cell entry into the CNS (17, 19), and rat T cells in the CNS of SCID mice appeared to satisfy this requirement independent of their specificity for Ag.

Although earlier events, including entry of T cells and BM-derived, CD11b-positive (MHC class II-negative and class II-positive) cells into the CNS, did not depend on the T cells’ specificity for myelin Ag, expression of MHC class II by a majority of the newly arrived CD11b-positive cells, and the subsequent development of inflammation and paralysis, did depend on T cell specificity for myelin. Thus, the results suggest the following order of Ag-independent and Ag-dependent events during the induction of CNS inflammation: 1) Ag-independent entry of activated T cells into the CNS; 2) Ag-independent, T cell-mediated recruitment of MHC class II-negative BM-derived cells into the CNS; 3) Ag-independent up-regulation of MHC class II expression and/or recruitment of MHC class II-positive cells; 4) Ag-dependent up-regulation of MHC class II expression and/or recruitment of MHC class II-positive cells; and 5) Ag-dependent inflammation and paralysis. To understand the mechanisms controlling susceptibility to and severity of T cell-mediated CNS inflammation, it will be necessary and should be possible to delineate the Ag-independent, cellular, and molecular interactions involving T cells and hemopoietic cells during the earliest stages of APC development, before the onset of clinical signs.

The current studies do not distinguish whether the role of BM-derived, CD11b-positive, myelomonocytic cells is restricted to Ag presentation or whether CNS pathology, inflammation, and/or paralysis are mediated by T cells, myelomonocytic cells, or both cell types in this model. However, using this xenogeneic cell-transfer approach, it should be possible to discern the cellular and molecular sources of paralytic activity by using T cell or BM donors possessing specific gene-targeted or other functional defects for selected regulatory, inflammatory, or toxic cell products. Furthermore, the Ab used in this study to detect BM-derived myelomonocytic cells reacts with multiple distinct subpopulations. To develop a more complete understanding of the cellular processes leading to CNS inflammation it will be necessary to define in more specific detail the nature and characteristics of the relevant subpopulations of BM-derived CD11b-positive cells.

In addition to CNS inflammation, BM-derived cells in the CNS may be involved in Alzheimer’s disease (43), CNS viral infection (44), tissue damage (45) or repair (46) following CNS trauma, and tumor pathogenesis (47, 48). Furthermore, controlled migration of BM-derived cells into the CNS may provide a strategy for introducing therapeutic agents or genes across the BBB (49). An understanding of mechanisms governing the development, trafficking, and ultimate functions of T cells and circulating BM-derived cells in the CNS may provide important insights into the regulatory processes that control the status of the CNS relative to a variety of diseases involving these cells. Using the interspecies transfer method in SCID mice, it should be possible to examine these processes in more detail.


   Acknowledgments
 
We thank Laura McGreevey for technical assistance with immunohistochemical staining.


   Footnotes
 
1 This work was supported by the U.S. Department of Veterans Affairs and the Nancy Davis Center Without Walls. Back

2 Address correspondence and reprint requests to Dr. Richard E. Jones, Research Service, P-3-R&D-23, Veterans’ Affairs Medical Center, 3710 S.W. U.S. Veterans Hospital Road, Portland, OR 97201. Back

3 Abbreviations used in this paper: MS, multiple sclerosis; BBB, blood-brain barrier; BM, bone marrow; BMC, BM cells; LN, lymph node(s); BP, myelin basic protein; EAE, experimental autoimmune encephalomyelitis; PPD, purified protein derivative of M. tuberculosis; SC, spinal cord. Back

Received for publication July 11, 2000. Accepted for publication October 6, 2000.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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