Virus-Cell Interactions

Preferential Small Intestine Homing and Persistence of CD8 T Cells in Rhesus Macaques Achieved by Molecularly Engineered Expression of CCR9 and Reduced Ex Vivo Manipulation

Matthew T. Trivett, James D. Burke, Claire Deleage, Lori V. Coren, Brenna J. Hill, Sumiti Jain, Eugene V. Barsov, Matthew W. Breed, Joshua A. Kramer, Gregory Q. Del Prete, Jeffrey D. Lifson, Adrienne E. Swanstrom, David E. Ott
Guido Silvestri, Editor
DOI: 10.1128/JVI.00896-19

ABSTRACT

Adoptive cell transfer (ACT) is a powerful experimental approach to directly study T-cell-mediated immunity in vivo. In the rhesus macaque AIDS virus model, infusing simian immunodeficiency virus (SIV)-infected animals with CD8 T cells engineered to express anti-SIV T-cell receptor specificities enables direct experimentation to better understand antiviral T-cell immunity in vivo. Limiting factors in ACT experiments include suboptimal trafficking to, and poor persistence in, the secondary lymphoid tissues targeted by AIDS viruses. Previously, we redirected CD8 T cells to B-cell follicles by ectopic expression of the CXCR5 homing protein. Here, we modify peripheral blood mononuclear cell (PBMC)-derived CD8 T cells to express the CCR9 chemokine receptor, which induces preferential homing of the engineered cells to the small intestine, a site of intense early AIDS virus replication and pathology in rhesus macaques. Additionally, we increase in vivo persistence and overall systemic distribution of infused CD8 T cells, especially in secondary lymphoid tissues, by minimizing ex vivo culture/manipulation, thereby avoiding the loss of CD28+/CD95+ central memory T cells by differentiation in culture. These proof-of-principle results establish the feasibility of preferentially localizing PBMC-derived CD8 T cells to the small intestine and enables the direct experimental ACT-based assessment of the potential role of the quality and timing of effective antiviral CD8 T-cell responses to inhibit viral infection and subsequent replication in small intestine CD4 T cells. More broadly, these results support the engineered expression of homing proteins to direct CD8 T cells to target tissues as a means for both experimental and potential therapeutic advances in T-cell immunotherapies, including cancer.

IMPORTANCE Adoptive cell transfer (ACT) of T cells engineered with antigen-specific effector properties can deliver targeted immune responses against malignancies and infectious diseases. Current T-cell-based therapeutic ACT relies on circulatory distribution to deliver engineered T cells to their targets, an approach which has proven effective for some leukemias but provided only limited efficacy against solid tumors. Here, engineered expression of the CCR9 homing receptor redirected CD8 T cells to the small intestine in rhesus macaque ACT experiments. Targeted homing of engineered T-cell immunotherapies holds promise to increase the effectiveness of adoptively transferred cells in both experimental and clinical settings.

INTRODUCTION

Effective cellular immune responses require the right cells, i.e., cells with the right specificities and effector activities, to be at the right place at the right time. Adoptive cell transfer (ACT) of genetically modified autologous T cells into mice, monkeys, and humans is a powerful experimental approach to modulate in vivo immune processes. Currently, adoptive T-cell immunotherapies that infuse patients with autologous T cells engineered ex vivo to express chimeric antigen receptors, CAR T cells, have shown efficacy against hematologic tumors which are readily accessed by the circulatory system, yet this approach has been less successful for solid tumors (1–3). Therefore, the ability to direct localization of infused engineered cells by using homing proteins has the potential to improve the efficacy of ACT for tissue-localized anticancer and antiviral targets (4–9).

The failure of T-cell responses to completely control or clear AIDS virus infection is attributed in part to the differential kinetics of exponential viral replication early after infection versus the lagging development of the initial T-cell response, which is further hindered by virally induced depletion of CD4 T cells that otherwise would contribute to the development of optimal cellular immune responses. This head start allows the virus to become widely disseminated, damaging the immune system and establishing long-lived viral reservoirs before the nascent T-cell response begins to suppress viral replication or clear infected cells. Even with conventional vaccines that induce antiviral memory T-cell responses, it appears that the postinfection delay for anamnestic expansion of vaccine-induced T cells, ineffective differentiation into potent cytotoxic T cells, and suboptimal trafficking of effector T cells to sites of viral replication limits the effectiveness of viral control. ACT using large numbers of CD8 T cells engineered to express AIDS virus-specific T-cell receptors (TCRs) that place effective antiviral T-cell immunity in AIDS virus-targeted tissues, around the time of viral inoculation, provides a means to test this “too little, too late” theory of why cellular immunity is of limited effectiveness in controlling AIDS virus infection (10).

Recently, we demonstrated that infusing simian immunodeficiency virus (SIV)-specific TCR-engineered T cells 3 days after a high-dose intrarectal SIV inoculation reduced the number of founder viruses transmitted to rhesus macaques (11), establishing an initial demonstration of the potential of ACT with antiviral engineered T cells to alter AIDS virus infection.

In addition to timing, colocalization of antiviral T cells with their targets is required for T-cell recognition and function (9). Because most ACT experiments rely on the natural movement of T cells between blood and tissue for infused cell distribution (12, 13), there potentially are certain tissue sites into which most CD8 T cells fail to effectively enter due to a lack of chemotactic signaling, resulting in de facto viral sanctuaries, even in individuals where CD8 T-cell responses appear capable of controlling viral replication in other more accessible sites. B-cell follicles residing in secondary lymphoid tissues represent such an immune-privileged sanctuary site for residual viral replication and virus production, even in the setting of protective major histocompatibility complex allele-associated spontaneous “elite control” or during combination antiretroviral therapy (14–24). Our recent proof-of-principle demonstration of directed trafficking of bulk CD8 T cells to B-cell follicles in rhesus macaques through engineered ectopic expression of CXCR5 (25) is now being exploited to evaluate directed localization of engineered antiviral responses into this normally immune-privileged viral sanctuary to target and suppress SIV-infected CD4 T cells (D. E. Ott, unpublished data).

Gut-associated lymphoid tissue is an important target for extensive early AIDS virus replication, resulting in a massive depletion of resident CD4 T cells and subsequent compromise of intestinal epithelial barrier integrity and leading to microbial translocation into gut tissue, which induces chronic pathological systemic inflammation in the gut (26–30). ACT offers a promising approach to experimentally test ways in which effective antiviral CD8 T cells in the small intestine at the time of viral inoculation could prevent or attenuate viral replication and CD4 T-cell depletion. Infusing large numbers of CD8 T cells engineered for expression of both SIV-specific TCRs for antiviral function and a homing receptor for enhanced localization to the small intestine would place timely, defined, effective antiviral responses either before or just after infection to combat early infection in this highly susceptible tissue.

The C-C chemokine receptor 9 (CCR9 or CDw199), a member of the large G-protein-coupled chemokine receptor (GPCR) family, has been implicated in the homing of T cells to the small intestine by chemotaxis in response to its cognate ligand, the chemokine CCL25, which is produced primarily by the epithelial cells of the small intestine (31–37). In the digestive tract, the expression of CCL25 is tightly restricted to the small intestine and absent from neighboring tissues, including stomach and colon (35, 36, 38), with CCL25 concentrations being highest in the duodenum and falling in a gradient to the distal ileum region (39). Additional evidence for CCR9 acting as a homing protein is its expression in cancers that metastasize to the small intestine (40) along with studies from ccr9 transgenic knockout mice (41). CCR9 does not appear to be strictly required for homing to small intestine: experiments in mice demonstrate both CCR9-dependent and -independent trafficking to the small intestine (39, 42).

To establish whether CCR9 can function as a T-cell homing protein capable of localizing infused CD8 T cells to the small intestine in rhesus macaques, we engineered bulk peripheral blood mononuclear cell (PBMC)-derived CD8 T cells to ectopically express CCR9, using improved gene transduction and cell expansion procedures that enhance in vivo persistence. Adoptively transferred CCR9-expressing CD8 T cells showed preferential homing to the small intestine and extended persistence, establishing CCR9 as a bona fide homing protein in primates and opening up new ways to study the role of the small intestine in AIDS virus immunology and pathogenesis.

RESULTS

To express CCR9 in T cells, we designed a retroviral vector that expresses the rhesus macaque CCR9 gene. However, GPCRs contain a serine-rich region in the C terminus of their cytoplasmic tail that binds arrestin and directs its rapid endocytosis (43–46). For instance, inherited gene mutations in humans which truncate the serine-rich region in the C terminus of CXCR4 GPCR result in loss of receptor endocytosis and are associated with WHIM syndrome, a dominant gain-of-function immunological disease caused by the dysregulation of CXCR4 signaling (47, 48). While arrestin-mediated endocytosis has not been formally demonstrated for CCR9, based on this conserved GPCR property, we truncated the C terminus of CCR9 in our vector to potentially stabilize and increase its surface expression, placing a premature stop codon in place of tryptophan at codon 343 and thereby removing the amino acids just beyond the 8th alpha helix, which includes the serine-rich region (49). Transduction of PBMC-derived primary CD8 T cells with this CCR9 vector induced ectopic cell surface expression of CCR9 (Fig. 1A).

FIG 1
FIG 1

CCR9 transduction of primary rhesus macaque T cells confers functional CCL25-mediated signaling and chemotaxis. Analyses of CCR9-transduced CD8 T cells are presented. (A) A dot plot of CCR9/CD28 flow cytometry analysis of a CCR9 vector-transduced primary T cell culture. (B) Near-infrared LI-COR ERK1/2 and p-ERK immunoblots of cell lysates, with minutes of CCL25 exposure denoted above each sample, molecular mass standards indicated at the left, and bands identified at the right. Total ERK detected varied by no more than 12% between samples in any blot. (C) Graph of the kinetics of pERK induction for each of the three independent experiments is plotted as the response relative to the maximal p-ERK value for each experiment. (D) Graph of specific migration by primary CD8 T cells transduced with our CCR9 vector in the presence of CCL25. Error bars indicate standard deviations.

In vitro functional evaluation of CD8 T cells transduced with CCR9.To confirm the ability of the CCR9 vector to confer ligand-specific signaling, we examined CCL25-mediated signaling in primary rhesus macaque CCR9 CD8 T-cell transductants by quantitative near-infrared immunoblot analysis for induced phosphorylation of the ERK1 and ERK2 (pERK1/2) protein kinases, a key step in the GPCR signaling cascade (50). Serum-starved CCR9 CD8 T-cell cultures were stimulated with CCL25, and cell lysates from several time points were analyzed (Fig. 1B). The results from three independent experiments showed rapid induction of pERK1/2 in the presence of CCL25, which peaked at 5 to 15 min before decreasing to a somewhat lower level (Fig. 1B and C). In contrast, the matching unmodified CD8 T cells failed to generate any detectable pERK1/2 in the presence of CCL25 (Fig. 1B), consistent with ligand-specific signaling in the CCR9 transductants.

To determine whether the CCR9 vector could confer chemotactic migration activity to T cells, we transduced primary rhesus macaque CD8 T cells with the CCR9 vector and examined the ability of the CCR9 transductants to migrate in response to CCL25 in a transwell assay (Fig. 1D). CCR9 transductants specifically migrated in response to CCL25, with more cells trafficking across the transwell barrier in the presence of CCL25 than in its absence (Fig. 1D), demonstrating that transduction of T cells with the CCR9 vector enables CCL25-dependent chemotaxis to primary CD8 T cells in vitro. These results, taken together with the signaling data (Fig. 1B and C), demonstrate that the CCR9 vector successfully confers specific CCL25-dependent functions in vivo.

Improving cell quality for rhesus macaque adoptive transfer experiments.In addition to tissue-specific homing, two additional factors for successful tissue-specific antiviral ACT are the overall ability of the infused virus-specific engineered T cells to be effectively distributed to the relevant virus-infected tissues to effect chemokine-specific homing, especially secondary lymphoid tissues, and, once present, to persist in those tissues. Our previous ACT experiments relied on extensive expansion of a small number of CD8 T cells, either from a single native SIV-specific CD8 T-cell clone isolated from PBMC (51, 52) or ∼50,000 CD8 T-cell transductants (11) to a final infusion product of 1 × 109 to 8 × 109 cells, a process which typically requires ∼12 weeks of culture with anti-CD3 stimulation every 2 weeks (11, 51, 52). However, upon infusion these extensively cultured cells exhibit marginal, if any, persistence and poor distribution to lymphoid tissues with predominant localization to the lung mucosa, as assessed by bronchoalveolar lavage sampling (11, 51–54, and D. E. Ott, unpublished data).

To understand the impact that time and successive stimulation in culture have on the transduced T cells, we performed preliminary ACT experiments using T cells expanded in culture for different periods of time. Consistent with accumulating experience in the field (55–61), the results suggested that shorter expansion times, with fewer stimulations, provided better persistence and distribution in secondary lymphoid tissues of the infused cells (data not shown). Phenotypic analyses of these preliminary experiments suggested that improved overall tissue distribution and persistence of infused CD8 T cells was associated with the subset expressing the CD28+/CD95+ memory differentiation phenotype, which can be used to define central memory T cells (Tcm) in the rhesus macaque system (62).

To better assess the relationship between a CD28+/CD95+ Tcm phenotype and cell distribution and persistence following infusion, CD8 T cells isolated from five sequential blood collections spanning 10 to 60 days preinfusion from rhesus macaque A (RM-A) were each individually stimulated with nonhuman primate anti-CD2, -CD3, and -CD28 paramagnetic beads immediately after isolation and then transduced 2 days later, using a retroviral vector with an irrelevant insert. Each of the five cell cultures was subsequently restimulated in the respective expansion cultures every 2 weeks, using anti-CD3 and lethally irradiated human PBMC as feeder cells (63) until infusion, as previously described (52).

A preinfusion flow cytometry analysis of the cultures for Tcm (CD28+/CD95+) revealed that, of the five sequential cultures, only the T cells cultured for 10 days with a bead stimulation followed by a single anti-CD3/feeder cell stimulation had a high frequency (43%) of CD28+/CD95+ CD8 Tcm cells (Fig. 2A). The remaining 57% of CD8 T cells in the 10-day culture exhibited the CD28−/CD95+ T cell effector memory (Tem) phenotype (62). Overall, the 10-day culture had a total proportion of CD28+ T cells similar to that of freshly isolated CD8 T cells from the animal: 49% total CD28+ CD8 T cells, consisting of 24% CD28+/CD95+ and 25% CD28+/CD95− (naive T cells), versus 51% CD28−/CD95+ (Fig. 2A, far right). In the four 17- to 60-day cultures, the proportion of CD28+/CD95+ cells rapidly decreased with the duration of culture and number of anti-CD3/feeder cell stimulations, consistent with ex vivo differentiation of Tcm to Tem (Fig. 2A).

FIG 2
FIG 2

Tissue distribution and persistence of infused T cells in animal RM-A is related to maintenance of a CD28+, Tcm phenotype. Dot plots of flow cytometry CD28/CD95 analysis gated for infused CD8 T cells (CD3+/CD8+/CTV+) in animal RM-A samples are presented. (A) Five sequentially isolated and transduced CD8 T-cell cultures expanded ex vivo for 10 to 62 days postisolation and a freshly isolated, unmanipulated PBMC culture with days in culture before infusion presented above each plot, with total of CD28+ CD8 T cells in the freshly isolated culture highlighted in blue. (B) Early kinetics of the levels of infused cells in PBMC samples collected between 15 min and 4 days postinfusion. (C) Long-term persistence of infused cells in PBMC, peripheral lymph node, and BAL collections from days 2, 7, and 35 postinfusion.

To examine tissue distribution and persistence in vivo, the five RM-A cell cultures, each carried for a different duration, were combined to assess the influence of time in culture and CD28 expression on infused cell distribution and persistence. The mixture, containing a total of 2 × 109 cells, was labeled with CellTrace Violet (CTV) for in vivo tracking, and the resulting infusion product was infused intravenously into RM-A. Even though most of the cells in the infusion product had the Tem phenotype (82% CD28−/CD95+) and only 18% bore the Tcm, CD28+/CD95+ phenotype, the infused (CTV+) T cells present in PBMC sampled at just 15 min postinfusion had nearly equal proportions of Tcm and Tem phenotypes, 42% CD28+/CD95+ and 57% CD28−/CD95+, respectively (Fig. 2B). Considering that nearly all of the infused CD28+/CD95+ cells came from the twice-stimulated, 10-day culture (Fig. 2B), the rapid decrease of infused CD28−/CD95+ cells in the PBMC compartment apparently reflects the disappearance of a large portion of the more extensively cultured cells, similar to the rapid loss of extensively cultured infused T cells from circulation observed in our prior ACT experiments (11, 51, 52, and unpublished results). The decline in frequency of infused Tem phenotype cells in PBMC continued until day 4 postinfusion, when they represented less than 8% of circulating CTV+ cells despite initially making up more than 80% of the infusion product (Fig. 2B).

To examine the tissue distribution and persistence of the infused cells, we compared the long-term presence of infused CD8 T cells in peripheral lymph node and bronchoalveolar lavage fluid (BAL) samples to those in PBMC at days 2, 7, and 35 postinfusion (Fig. 2C, left). Bronchoalveolar airways are a conveniently accessed mucosal site that serves as a proxy measure for lung localization, a site to which infused cells have localized to and persisted in our prior ACT experiments (11, 51, 52, and unpublished data). The level of CD28+/CD95+ CD8 T cells in the PBMC samples was essentially maintained at or above the 84% 2-day level through the 35-day postinfusion analysis (Fig. 2C, top row). Similarly, nearly all of the CTV+ infused cells identified in the peripheral lymph node samples had a Tcm phenotype (97 to 98%) from day 2 through 35 sampling (Fig. 2C, middle row), with few Tem cells present (<2% CTV+/CD28−/CD95+). The overwhelming levels of Tcm and scarcity of Tem in lymph nodes are consistent with our previous results showing a failure of Tem CD8 T cells to enter and/or persist in these secondary lymphoid tissues in our prior ACT experiments using extensively cultured CD8 Tem cells (11, 51, 52, and unpublished data). Thus, CD8 Tcm are superior for targeting lymph nodes and potentially other secondary lymphoid organs.

In contrast to the lymph node samples, the vast majority of CTV+ cells in the BAL samples were Tem cells (74 to 88% CD28−/CD95+) throughout the 35-day sampling, with only 12 to 26% showing the CD28+/CD95+ Tcm phenotype, consistent with preferential localization of Tem cells to the lung (11, 52). However, regardless of the CD28 expression status, the frequencies of the infused cells decreased between the 7-day and 35-day time points in all of the sample sets, suggesting that the Tem cells can persist in the lung (Fig. 2C, right). Taken together, these results suggest that better persistence in PBMC and overall lymphoid tissue distribution is achieved by maintaining CD28+ expression on the cells during ex vivo manipulation and culture. This appears to be due in part to less time in culture and fewer stimulation cycles, consistent with abbreviated ex vivo human T-cell expansion culture, resulting in improved T-cell function in ACT experiments in human tumor xenographed NOD-SCID mice (55).

Engineering central memory CCR9-transduced CD8 T cells for small intestine homing and persistence in rhesus macaques.To examine the potential of engineered CCR9 expression to direct CD8 T cells into the small intestine, we transduced freshly isolated bulk PBMC with the CCR9 vector. To minimize the length of time in culture and the number of stimulation cycles yet still produce sufficient quantities of engineered T cells for infusion, we devised a rapid large-scale transduction/expansion procedure that produces ∼2 × 109 to 16 × 109 engineered rhesus macaque CD8 T cells in 14 days with only two CD3 antibody-based stimulations. To examine the potential for engineered CCR9-mediated homing to the small intestine, rhesus macaque 1 (RM-1) underwent leukapheresis, yielding ∼2 × 108 PBMC that were immediately stimulated with nonhuman primate anti-CD2, -CD3, and -CD28 paramagnetic beads and transduced with the CCR9 vector 2 days postisolation/poststimulation in a large-scale transduction. At 1 week postleukapheresis, the transduced culture was restimulated by coculture with CD3 and CD28 antibodies and lethally irradiated human PBMC feeder cells and then infused 1 week later. This 2-week transduction and expansion procedure yielded 2.7 × 109 CD8 T cells, which were CTV labeled before infusion. Flow cytometry of the infusion product confirmed that 41% of the infused cells were CCR9 transductants, i.e., CTV+/CCR9+. The infusion product contained 12% Tcm CD28+/CD95+ and 29% CCR9+ Tem CD28−/CD95+ phenotypes (Fig. 3A, CD95+ data not shown), which is a considerably lower proportion of Tcm than that of the endogenous PBMC. Since ex vivo-stimulated T cells constitutively express CD95, the positive status of CD95 expression of the infused T cells, although monitored, is not included throughout the rest of this report. To delineate between the transduced and untransduced CD8 T cells in the transduced CTV+ infusion product and postinfusion flow cytometry samples, CD8CCR9 denotes the CTV+/CCR9+ CD8 T-cell transductants and CD8UT identifies the untransduced CTV+/CCR9− CD8 T cells in the infusion product that were exposed to the CCR9 retroviral vector yet failed to be transduced and were subsequently labeled with CTV.

FIG 3
FIG 3

Analysis of the presence of infused CD8CCR9 and Tcm phenotypes in animal RM-1 early infusion samples. Dot plots of flow cytometry analyses of CD8 T cells (CD3+/CD8+) in PBMC from animal RM-1 are presented. (A and B) The CD28 and CCR9 status of CD8 T cells in infusion product, infused CD8 T cell (CTV+) at 15 min and 24 h postinfusion, and endogenous (CTV−) samples (A) and infused CD8 T cells (CTV+) as a percentage of all CD8 T cells at 15 min and 24 h (B). Gating for the infused, endogenous, and all CD8 T-cell populations is indicated above their respective plots.

Analysis of infused CD8CCR9 in PBMC and lung mucosa.Flow cytometry CTV, CCR9, and CD28 analysis of a 15-min postinfusion PBMC sample found that the total infused CD8 T cells, CD8CCR9 plus CD8UT, accounted for 22% of total circulating CD8 T cells (Fig. 3B, left plot), with 69% of the infused cells being CD28+ (Fig. 3A, second plot), a rapid increase in frequency of CD28+ T cells over the 35% Tcm cells present in the infusion product (Fig. 3A, first plot). At 24 h postinfusion, the frequency of total infused cells increased to 43% in the 24-h PBMC sample (Fig. 3B, right plot), with 62% of those cells being CD28+ (Fig. 3A, third plot). These results are a great improvement of infused cell persistence in PBMC due to the presence of Tcm phenotypic CD8 T cells compared to levels in our prior ACT experiments (Fig. 2) (11, 52, and unpublished data.)

To examine the tissue localization of infused CCR9 transductants, animal RM-1 was euthanized and necropsied 2 days postinfusion, and the presence of both CD8CCR9 and CD8UT cells and their corresponding frequencies of Tcm phenotype in PBMC, BAL fluid, and other relevant tissues were determined by flow cytometry analysis of mononuclear cell suspensions for CTV+/CCR9+/CD28+ CD8 T cells. Because the infusion product was a mix of 41% CD8CCR9 and 59% CD8UT cells (Fig. 3A), the untransduced CD8UT cells serve as an internal control for nonspecific homing; thus, comparing the relative frequencies of CCR9+ and CCR9− cells within the CTV+ population indicates preferential homing.

The initially observed frequencies of infused CD8 T cells in PBMC were maintained after 2 days (Fig. 4), with 40% of the CD8 T cells in the PBMC sample being infused CTV+ CD8 T cells, of which 32% were CD8CCR9 cells and 62% expressed CD28, a level similar to the frequency of CD28 expression on endogenous CTV−/CD3+ T cells (69%). The day 2 prenecropsy BAL samples also contained high frequencies of the infused cells (Fig. 4), with 55% of all CD8 T cells in the BAL sample being infused cells; however, unlike the PBMC samples, a large majority of the infused CD8 T cells in BAL samples were CD28− (92%), consistent with Tem trafficking to mucosal and peripheral nonlymphoid sites (64), a result similar to that of our prior adoptive T-cell transfers (11, 51, 52) and the data depicted in Fig. 2. Furthermore, the frequency of CD8CCR9 cells in the BAL fluid was 22%, with most being CD28−, a decline from the 41% in the infusion product (Fig. 3A), suggesting that they avoided this site.

FIG 4
FIG 4

Analysis of both infused and endogenous CD8 T cells in animal RM-1 PBMC and BAL necropsy samples at 2 days postinfusion. Dot plots of flow cytometry analyses of CD8 T cells in PBMC and BAL necropsy samples from animal RM-1 are presented. Gating for all CD8 T-cell, infused, and endogenous populations is indicated above their respective plots. Gates for CCR9 and CD28 in the infused and endogenous analyses were set using the PBMC endogenous plot (top right) as a standard.

Localization of infused CD8CCR9 cells to small intestine.Of the gut tissue necropsy samples, those from the duodenum clearly stood out, with infused cells (CTV+) making up 6% of the total duodenal CD8 T-cell population by flow cytometry (Fig. 5, top row). Flow cytometry CCR9/CD28 analysis of the CTV+ CD8 T-cell population in duodenal samples demonstrated a 60% frequency of CD8CCR9, an increase over the 41% CD8CCR9 present in the infusion product (Fig. 3A). Given the 6% overall frequency of the infused cells, CD8CCR9 cells made up 4% of the total CD8 T cells in the duodenal sample. Also, 42% of the infused cells were CD28-expressing CD8CCR9 cells. In contrast, the levels of CCR9+ and CD28+ in the CD8+/CTV− endogenous duodenum population were lower, with 31% of this population being CCR9+ and 23% being CCR9+/CD28+ (Fig. 5).

FIG 5
FIG 5

Analysis of both infused and endogenous CD8 T cells in animal RM-1 gut tissue necropsy samples at 2 days postinfusion. Dot plots of flow cytometry analyses of CD8 T cells in gut tissue necropsy samples from animal RM-1 are presented. Gating for all CD8 T-cell, infused, and endogenous populations is indicated above their respective plots. Gates for CCR9 and CD28 in the infused and endogenous analyses were set using the PBMC endogenous plot shown in Fig. 4, top right, as a standard.

Examining the distribution of the infused cells in the small intestine by confocal fluorescence microscopy revealed CTV+ T cells present throughout the gut sample (Fig. 6A) in both the lamina propria and lymphoid aggregate regions (Fig. 6A and B), demonstrating that the CD8CCR9 T cells distribute throughout both mucosal and lymphoid tissues in the small intestine.

FIG 6
FIG 6

Visualization of infused cells in the small intestine of animal RM-1. Confocal fluorescence microscopy detecting the CTV fluorescent infused cells (pseudocolored blue) in small intestine are presented. (A) Overview with the border of the lymphoid aggregate, LA, and lamina propria, LP, tissues highlighted with a dot-dashed white line. (B) Closeup examples of lamina propria and lymphoid aggregate tissue sections. All scale bars represent 100 μm.

Sampling of the cecum detected considerably fewer total CD8 T cells, with only 3,500 CD8 T-cell-gated events captured, with 1% of those cells being infused cells (CTV+). However, within the infused cell population, 51% were CD8CCR9 cells and 44% were CCR9+/CD28+ (Fig. 5, second row). In stark contrast, colon samples, containing 1% infused CD8 T cells, were nearly devoid of CD8CCR9 cells, with 96% of those CTV+ CD8 T cells identified being CD8UT cells that were nearly exclusively CD28+. Accordingly, the native population exhibited a profile very similar to that of the infused CD8 T-cell population, nearly all CCR9−/CD28+ (Fig. 5, third row). Finally, the few infused cells found in sampling of the rectum (1%) also were almost exclusively CD8UT with few, if any, CD8CCR9 cells, frequencies matching the corresponding endogenous CD8 cells in the tissue. Unlike the colon, about half of the few infused cells present were CD28−, while the endogenous cells were even more skewed to the effector memory phenotype, 63% CD28− (Fig. 5, bottom row).

Taken together, these data show that both endogenous CCR9+ and the infused CD8 T cells engineered for expression of CCR9 preferentially localize in the small intestine, especially the upper small intestine (sampled here as duodenum), and appear to be relatively excluded from the colon and rectum, consistent with the reported restricted expression of CCL25 to the small intestine (35, 36, 38, 39). Furthermore, infused cells found in all of the gut tissues exhibited an enrichment of those with a CD28+ central memory phenotype over the infusion product, mostly mirroring CD8 expression in their endogenous CD8 T-cell population counterparts.

Presence of CD8CCR9 cells in lymph nodes.To investigate the presence of infused CD8CCR9 cells in secondary lymphoid tissues, we examined mesenteric, inguinal, and axillary lymph node samples for the presence of infused cells. Mesenteric lymph nodes drain lymph from the intestinal tract, and cells destined to become small intestine homing T cells are primed in these lymph nodes, acquiring expression of the α4β7 integrin and CCR9 by exposure to retinoic acid produced by mesenteric lymph node stromal cells. These primed tissue-targeted T cells then travel through the efferent lymph, eventually returning to and entering the small intestine via the blood (65–68). CTV+ CD8 T cells collected from the mesenteric lymph node sample at necropsy consisted of 24% CD8CCR9 cells, considerably lower levels than those seen in the duodenum tissue samples, with most of these cells being CD28+, as expected for lymph node residency (Fig. 7, top row). However, even though the endogenous CD8 T cells in the sample were nearly all CD28+, there were few, if any, CCR9-expressing CD8 T cells, consistent with the current gut priming/homing model (65–68) and the reported lack of CCR9+ CD8 T cells in murine mesenteric lymph nodes (69).

FIG 7
FIG 7

Analysis of both infused and endogenous CD8 T cells in animal RM-1 lymph node necropsy samples at 2 days postinfusion. Dot plots of flow cytometry analyses of CD8 T cells in lymph node necropsy samples from animal RM-1 are presented. Gating for all CD8 T-cell, infused, and endogenous populations is indicated above their respective plots. Gates for CCR9 and CD28 in the infused and endogenous analyses were set using the PBMC endogenous plot shown in Fig. 4, top right, as a standard.

In contrast, flow cytometry analysis of axillary and inguinal lymph nodes showed a barely detectable fraction of CD8CCR9 cells in these tissues, with ∼94% of the infused cells detected being the CD8UT component with the CD28+ Tcm phenotype, again matching the profile of the endogenous CD8 T cells in these samples (Fig. 7, middle and bottom rows). Taken together, CCR9 expression on infused cells produces a distinct homing pattern, favoring small intestine and its draining lymph nodes and seemingly avoiding peripheral lymph nodes.

Long-term homing and persistence of CD8CCR9 cells.To assess the ability of infused CD8CCR9 cells to home to and persist in the small intestine over time, we carried out an ACT experiment followed by long-term sampling using rhesus macaque 2 (RM-2). For this experiment, we produced two T-cell cultures: a CTV-labeled CCR9-transduced culture containing both CD8CCR9 and CD8UT T cells and, as an unambiguous negative control, a CellTrace Far Red dye (CTFR)-labeled unmodified CD8 T-cell culture, using the same leukapheresis and rapid T-cell molecular modification and expansion procedure as that used for RM-1. The CCR9 transduction culture contained 2.4 × 109 CTV+ T cells composed of 53% CD8CCR9 transductants, with 12% of the culture expressing both CCR9 and CD28, a lower level than that of fresh endogenous CD8+/CD28+ T cells (Fig. 8A and B). The parallel unmodified culture consisted of 1.2 × 109 CTFR+ cells with only a few dimly staining endogenous CCR9+ T cells present. Thirty-nine percent of the CD8 T cells in the unmodified culture expressed CD28 (Fig. 8C), a level similar to that of the endogenous sample (Fig. 8B). At 15 min postinfusion, the CTV+ CD8 T cells accounted for 7% and CTFR+ cells were 3% of the total CD8 T cells, frequencies which rose to 14% and 5%, respectively, at 24 h (Fig. 8D). The proportion of CD28+ CD8CCR9 in the CTV+ fraction rose steadily from the infusion product (12%) through the 24-h PBMC sample (28%) (Fig. 8A). In both the transduced (CTV+) and unmodified (CTFR+) populations, the proportions of CD28-expressing cells increased over time relative to their respective infusion products (Fig. 8A and C).

FIG 8
FIG 8

Analysis of the presence of infused CD8CCR9 and Tcm phenotype in animal RM-2 early infusion samples. Dot plots of flow cytometry analyses of CD8 T cells in PBMC from animal RM-2 are presented. (A) CD28 and CCR9 status of the transduced CTV+ CD8 T-cell component of the infusion product and the 15-min and 24-h postinfusion PBMC samples. (B) CD28 and CCR9 status of CD8 T cells in the endogenous (CTV−) PBMC sample. (C) CD28 and CCR9 status of the unmodified CTFR+ CD8 T-cell component of the infusion product and the 15-min and 24-h postinfusion PBMC samples. (D) Frequencies of both CTV+ and CTFR+ infused cells in all CD8 T cells at 15 min and 24 h. Gating for the infused, endogenous, and all CD8 T-cell populations is indicated above their respective plots.

To assess the relative localization of infused CD8CCR9 T cells in various tissues between RM-1 and RM-2, we compared the frequencies of the CD8CCR9 cells in specific necropsy (RM-1) or biopsy (RM-2) tissue samples obtained at day 2 postinfusion. To account for differences in the numbers of CD8CCR9 T cells infused for RM-1 and RM-2, we normalized the percentage of the CD8CCR9 cells in the entire CD8 T-cell population in the 2-day samples observed in general circulation, i.e., PBMC, and of those that localized to tissues (duodenum, rectum, and peripheral lymph node) to the percentage of CD8CCR9 in their corresponding infusion products (Fig. 9). The resulting relative frequencies showed that among tissues, the infused CD8CCR9 T cells were highest in the duodenal samples, indicating strong localization to the small intestine. In contrast, rectal and lymph node samples in both animals had considerably lower levels of CD8CCR9 T cells and markedly less than that in general circulation, entirely consistent with preferential tissue localization of CD8CCR9 T cells infused to the small intestine.

FIG 9
FIG 9

Normalized CD8CCR9/CTV+ T-cell frequencies in tissue necropsy and biopsy samples from day 2 postinfusion in animals RM-1 and RM-2, respectively. A graph of flow cytometry analysis frequencies of CD8CCR9 cells in samples from animals RM-1 and RM-2 normalized to their respective frequencies in their corresponding infusion products is presented for PBMC, duodenum, rectum, and peripheral lymph node samples.

Preferential persistence of CD8CCR9 T cells in the small intestine.The presence of the infused CD8CCR9 cells in longitudinal biopsy tissue samples from RM-2 remained higher in the duodenal biopsies than the rectal biopsies and peripheral lymph node samples over 48 days of sampling (Fig. 10A), demonstrating that the infused CD8CCR9 cells preferentially localized to the small intestine, unlike other tissues, and persisted there. Due to the invasive nature of excisional lymph node biopsies and the low frequencies of CD8CCR9 cells in these tissues, peripheral lymph nodes were only biopsied at days 2 and 9 postinfusion.

FIG 10
FIG 10

Persistence of infused cells in animal RM-2 tissue samples. (A) Graphs of flow cytometry analyses are presented for percentage of CD8CCR9 cells of all CD8 T cells in duodenal, rectal, and peripheral lymph node samples between 2 and 102 days postinfusion. Due to animal protocol limitations, lymph nodes were only sampled at days 2 and 9 postinfection. (B) Graphs of flow cytometry analyses are presented for percentage of CD8CCR9 CD28+ transduced (CTV+) cells in duodenal, rectal, and peripheral lymph node samples between 2 and 102 days postinfusion. (C) Graphs of flow cytometry analyses for percentage of CTV+/CD8CCR9 and CTFR+ T cells of total CD8 T cells show differential persistence of CTV+/CD8CCR9 and CTFR+ (unmodified CD8 T cells) in duodenal samples between 2 and 102 days postinfusion. ND, not done; *, too few T cells in the pinch biopsy sample for analysis.

The CD28+ status of the CD8CCR9 cells in tissues mostly mirrored that of RM-1 (Fig. 10B). Nearly half of the CD8CCR9 cells in the duodenum samples were CD28+, and almost all of these cells in the lymph node were CD28+ (Fig. 10B). The rectum samples had widely varying CD28+ frequencies in their CD8CCR9 populations, which, on average, mirrored the nearly 50% CD28+ frequency observed in RM-1.

To address the possibility that the apparent preferential localization of the CD8CCR9 T cells to the small intestine was somehow an artifact of the infusion procedure, we compared the relative presence of the infused CD8CCR9 cells with that of the CTFR+ unmodified CD8 T cells (Fig. 10C). For all time points, the duodenal biopsy samples contained higher frequencies of CD8CCR9 than the unmodified CTFR+ CD8 T cells despite nearly equal numbers of cells in the infusion product (1.3 × 109 CD8CCR9 versus 1.2 × 109 CTFR+) (Fig. 8A and C). Based on Fig. 10C, the spread between the respective frequencies (CTV+ versus CTFR−) increased after day 9, with CD8CCR9 being 60%, 80%, 95%, and 78% of the infused (CTV+) CD8 T-cell population analyzed at days 9, 23, 48, and 102, respectively, consistent with the enhanced accumulation/retention and persistence of CD8CCR9 T cells in the small intestine being due to ectopic CCR9 expression.

Overall persistence of infused cells in PBMC.The extended persistence of infused cells observed in the tissue samples, particularly the duodenal biopsies, was also observed in the PBMC samples (Fig. 11). The frequency of infused cells steadily decreased over time, with CTV+ and CTFR+ T cells representing 0.04% and 0.05% of the total CD8+ T cells, respectively, at day 148 postinfusion (Fig. 11A). The similarity of these frequencies indicates that CCR9 expression alone does not have any overt effect on persistence in general circulation per se. As expected, persistence in the PBMC compartment was highly related to CD28 expression, as evidenced by the increasing proportion of CD28+ cells within the persisting infused population in the PBMC over time (Fig. 11B). Even as the total proportions of infused cells decreased, the relative contribution from CD28+ cells increased, from 33% CD28+ in the infusion product to 77% at day 27 postinfusion and subsequent maintenance at or above that level through day 148. Thus, our rapid T-cell expansion protocol improved the overall persistence of our infused cells by retaining and maintaining more of the Tcm cells from the initial donor leukapheresis material.

FIG 11
FIG 11

Persistence of CCR9/CTV+ versus unmodified CTFR+ CD8 T cells and CD28 phenotyping in RM-2 PBMC samples. Graphs of flow cytometry data obtained from 15-min to 148-day postinfusion PBMC samples are presented, showing the percentage of infused CCR9+/CTV+ and CTFR+ CD8 T cells in all CD8 T cells (A) and the frequency of CD28 expression among the total infused (CCR9+/CTV+ and CTFR+) CD8 T cells (B).

DISCUSSION

Here, we demonstrate both preferential in vivo localization of PBMC-derived CD8 T cells to the small intestine by engineered ectopic expression of CCR9 and enhanced persistence achieved by maintaining Tcm-like cells in the infusion product. The ability of the transduced CD8CCR9 T cells to effectively localize to the small intestine compared to their coinfused unmodified counterparts directly confirms that the CCR9 chemokine receptor functions as a homing protein. Despite clear localization of the infused CD8CCR9 cells to small intestine, demonstrated in serial duodenal biopsies, these cells were present at considerably lower frequencies in downstream gut tissues, with colon and rectum essentially devoid of infused CD8CCR9 cells, consistent with the expression pattern of the CCL25 chemokine (35, 36, 38) and a dependence of CCR9-CCL25 interactions for small intestine homing.

Relatively few, if any, CD8CCR9 cells were present in the axillary or inguinal lymph node samples, mostly matching the absence of endogenous CCR9+ CD8 T cells in these tissues. In contrast, the mesenteric lymph node sample contained some of our CD8CCR9 transductants yet considerably fewer than the duodenal sample. The few endogenous CCR9+ CD8 T cells present in the mesenteric lymph node are consistent with data from mice showing that CD4 T cells are the predominant CCR9-expressing T-cell type in this tissue (69), presumably as helpers for T-cell priming from the draining gut lymph nodes (65–68). The presence of CD8CCR9 T cells in the mesenteric lymph node could be due to the higher levels of circulating CCR9 transductants that either drain from other gut tissues that fail to express CCL25 and, thus, do not support CCR9-mediated homing or simply fail to enter the small intestine, potentially due to a lack of expression of endothelial transmigration factors (discussed below).

While CCR9 expression is required for redirecting CD8 T cells into the small intestine, another key component of tissue homing involves transiting from the bloodstream into the interstitial space. CXCR3 expression has emerged as an important factor in homing because it induces T cells in circulation to extravasate into the interstitial space of various tissues and finally the lymphatic system, enabling T cells to detect and follow specific chemotactic homing signal gradients produced by chemokine-secreting cells that stimulate the corresponding T-cell chemokine receptor, in this case CCR9 (70–72).

Similarly, α4β7 integrin expressed on circulating cells, including endogenous CCR9+ CD8 T cells, can bind mucosal addressin cell adhesion molecule-1 (MadCAM-1), which is expressed in postcapillary venules present in the small intestinal lamina propria (39, 42, 73), immobilizing circulating CCR9+ CD8 T cells at this site and facilitating their ability to detect CCL25 and follow chemotactic gradients to cross the lamina propria. Thus, both CXCR3 and α4β7 integrin act as initiating factors for T-cell homing. The infusion products from both RM-1 and RM-2 contained a majority of cells with dual expression of α4β7 integrin and CXCR3 (∼60%) with even higher levels of cells expressing at least one of the two markers (80%), providing the potential for transendothelial/extravasation activity for CCR9 homing in most of the infused CD8CCR9 transductants (Fig. 12).

FIG 12
FIG 12

Flow cytometry analysis of α4β7 and CXCR3 expression. A dot plot of flow cytometry analysis showing α4β7 integrin and CXCR3 expression on the CTV+ CD8 T cells in the infusion product of RM-1 is presented.

Our modified ACT cell production protocol produced considerably better long-term persistence of detectable levels of infused cells in PBMC than in our past experiments, ≥148 days postinfusion versus 3 to 21 days observed in both published (11, 52) and unpublished work, involving cell preparations subjected to longer and more extensive ex vivo manipulations. Persistence in PBMC was tightly correlated with the preservation of Tcm CD28+/CD95+ phenotype and was not linked to expression of CCR9. Additionally, it appears that the traditionally defined Tcm phenotype comprises a spectrum of subtypes, some of which show superior postinfusion persistence (61). Our results are consistent with accumulating experience indicating that minimization of the duration and extent of ex vivo manipulations that preserve Tcm CD8 T cells in infusion products is associated with improved results in clinical ACT (55–61).

In addition to enhanced persistence, the more effective maintenance of CD28+ CD8 T cells during the engineering process also provided better overall cell distribution, with CD28 expression highly related to generalized secondary lymphoid tissue localization by the untransduced CD8 T cells, a pattern expected for Tcm cells (64). Indeed, for all tissues except for mesenteric lymph node, the expression pattern for CCR9 and CD28 by the infused cells essentially matched that of their endogenous CD8 T-cell counterparts in the same tissues, further indicating that our infused CD8 T cells mostly trafficked in a physiological manner.

While we focus on persistence in PBMC and lymphoid tissue, the disappearance of the CD28−/CD95+ Tem cells is not a priori due to their absence from the animal in toto. Tem cells typically are retained at mucosal and peripheral nonlymphoid sites, such as lungs and skin (64). The high frequency of Tem cells in BAL fluid (11, 51, 52), a readily accessed nonlymphoid mucosal tissue, supports the idea that the transition of CD8 T cells to a Tem phenotype upon extended culture may redirect them to nonlymphoid sites that are often not effectively surveyed in ACT experiments. For some experiments, e.g., suppressing the transmission of founder viruses in mucosal infection (11), infusion of Tem CD8 T cells could be advantageous.

Current follow-up work is probing the practical limits of preparation of engineered T cells for ACT, balancing minimization of ex vivo manipulations with cell culture expansion yields to optimize in vivo persistence, directed tissue localization, and effector activity. Sustained persistence of infused ex vivo engineered antiviral cells capable of achieving a functional cure or viral eradication provides an aspirational goal for these efforts.

The ability to target CD8 T cells to specific tissues has potential applications for anticancer adoptive T-cell immunotherapy. Targeting of adoptive T-cell immunotherapeutics to specific tissues using homing proteins could improve the efficacy of CAR T or TCR-transduced cytotoxic T cells by focusing the infused cells to tumor tissues. Some tumors express tissue-specific chemokines while other cancers, such as melanoma, metastasize to small intestine by expressing CCR9 (40, 74–78), phenomena that could be exploited to engineer CD8 T cells with both antitumor and tumor-homing properties.

Using homing proteins to localize CD8 T cells engineered with potent antiviral activities opens exciting possibilities to both improve our understanding of the parameters of effective T-cell-mediated control of AIDS virus infection and increase the potential efficacy of clinical T-cell immunotherapies. To date we have demonstrated proof of principle for three different approaches to preferential localization of infused cells to different tissues, using CCR7/CD62L for peripheral lymph node targeting (11) and CXCR5 for B-cell follicles (25 and this study), demonstrating the ability of CCR9 expression to localize infused cells to the small intestine. Current and follow-up studies engineering potent antigen-specific cytotoxic T cells with tissue-targeted homing are beginning to assess how best to apply these approaches for maximum experimental and potentially therapeutic benefit.

MATERIALS AND METHODS

Rhesus macaques.Three Indian rhesus macaques (Macaca mulatta) were studied. Rhesus macaque A (RM-A; internal designator T157), male, 5 years old, 5.7 kg, average of 1,481 CD8 T cells/μl blood preinfusion, was chronically infected with SIVmac239 but was positive for the protective Mamu A*01 and Mamu B*17 MHC alleles (unpublished data) and an elite controller, with plasma viremia between <15 and 2 × 102 SIV RNA copies/ml during the chronic phase of untreated infection. Rhesus macaque 1 (RM-1; internal designator ZC63), female, 12 years old, 7.9 kg, average of 496 CD8 T cells/μl blood preinfusion, had been inoculated intrarectally with SIVmac239X (79) but did not show virological or immunological evidence of infection. Rhesus macaque 2 (RM-2; internal designator 034), female, 5 years old, 6.4 kg, average of 1,036 CD8 T cells/μl blood preinfusion, was infected with SIVmac239X with persistent chronic-phase viremia in the range of 5 × 105 to 2 × 106 SIV RNA copies/ml of plasma. All three animals were housed at the National Cancer Institute, NIH, Bethesda, MD, and cared for in accordance with the Association for the Assessment and Accreditation of Laboratory Animal Care (AAALAC) standards in an AAALAC-accredited facility, and all procedures were performed according to protocols approved by the Institutional Animal Care and Use Committee of the National Cancer Institute (protocols AVP-022 and AVP-062; assurance number A4149-01). Work involving animals adhered to the standards of the Guide for the Care and Use of Laboratory Animals (80) in accordance with the Animal Welfare Act.

CCR9 vector construction and production.A modified rhesus macaque CCR9 gene was produced synthetically by GeneART (Invitrogen) and inserted into the modified MSGV1 vector between the NcoI and NotI restriction endonuclease sites (81). Retroviral vectors were generated by transfecting vector constructs into Phoenix-RD114 packaging cells (82).

Preparation and transduction of cells for adoptive transfer.For RM-A, PBMC were isolated from a 60-ml venous bleed and processed as described below. For RM-1 and RM-2, PBMC were collected by apheresis using a Spectra Optia apheresis system (TerumoBCT, Inc.) using the PBMC filter set and the pediatric tubing cartridge kit according to the manufacturer’s protocol. For both sources, PBMC were isolated from the blood/leukapheresis product by density centrifugation through Ficoll-Paque (GE Life-Sciences). PBMC were then resuspended in RPMI 1640 medium supplemented with 10% (vol/vol) fetal bovine serum (Gemini Bio Products), 10 mM HEPES buffer, 2 mM glutamine (Life Technologies), 50 IU/ml interleukin-2 (Peprotech), 100 IU/ml penicillin (Invitrogen), and 100 μg/ml streptomycin (Invitrogen) and stimulated with Miltenyi nonhuman primate T cell activation/expansion beads (consisting of anti-CD2, -CD3, and -CD28; number 130-092-919; Miltenyi Biotec) per the manufacturer’s recommendations. Sequential transductions were carried out on days 2 and 4 poststimulation with our truncated CCR9 expression vector, MSGV-CCR9, using Retronectin (TaKaRa Bio USA) as previously described (25). Transduced T-cell cultures were expanded for 7 days after PBMC collection and then restimulated with anti-CD3 (30 ng/ml; clone SP34-2; BD Biosciences) and anti-CD28 (30 ng/ml; clone CD28.1) in the presence of 40 million lethally irradiated human PBMC per 25 million T cells (52, 63, 83) as feeder cells. T cells from RM-1 and -2 were cultured for an additional 7 days and then infused. T cells from RM-A were cultured for 3 to 55 days after the second stimulation before infusion.

Flow cytometry.CD3 allophycocyanin-CY7 (SP34-2), CD45 V450 (HI30), CD4 Alexa-700 or BV605 (OKT4), CD8α phycoerythrin (PE)-CY7 (SK1), CD28 PE-CF594 (CD28.2), CD95 PE-CY5 or BV650 (DX2), and CCR9 fluorescein isothiocyanate or APC (112509) antibodies were obtained from BD Biosciences, and α4β7 antibody was obtained from the NIH Nonhuman Primate Reagent Resource. Flow cytometry was carried out essentially as previously described (84), with the acquisition of at least 100,000 CD8 T-cell events per analysis using either a BD LSRII or Fortessa flow cytometer (BD Biosciences). The gating strategy for CD8/CTV+, CD8/CTFR+, and endogenous CD8 T-cell populations is presented in Fig. 13. Complete blood cell counts were monitored using BD TruCount tubes (BD Biosciences) on a BD FACSVerse (BD Biosciences). Data analysis was performed using FCS Express 4 (De Novo Software).

FIG 13
FIG 13

Gating strategies of CD8 T cell populations. Flow cytometry dot plots of the gating strategy analyses used to identify the experimental CD8/CTV+, unmodified control CD8/CTFR+, and endogenous CD8 T cell populations examined in downstream analyses are presented.

ERK1/2 phosphorylation analysis.CCR9-mediated ERK1/2 signaling was examined by immunoblotting for total and phosphorylated ERK1/2 (pERK1/2) in CCL25-stimulated CD8 T-cell lysates essentially as previously described (25). Briefly, 1 × 106 cells were cultured at 37°C in serum-free medium for 18 h and then stimulated with 2 μg/ml of human CCL25 (PeproTech) with samples taken after 5, 10, and 15 min. Cell pellets were produced, lysed, and analyzed by quantitative near-infrared immunoblotting (85) using pERK1/2-specific rabbit antiserum (ab4819; Abcam) at a 1:1,000 dilution and an ERK1/2-specific mouse monoclonal antibody (66192-1-Ig; Proteintech Group, Inc.) at a 1:5,000 dilution, followed by washing and treatment with a two-color secondary antibody detection system using anti-mouse IRDye 680LT and anti-rabbit IRDye 800CW fluorescently labeled donkey secondary antibodies (LI-COR) and then analyzed, after washing, with an Odyssey infrared imaging system (LI-COR) using a laser intensity of between 1 and 5. Signal densities of bands were measured by the Odyssey 3.0 application software.

In vitro chemotaxis assay.Chemotaxis was measured as previously described (25). Briefly, using transwell inserts (5-μm pore size) on a 24-well plate (Costar), the upper and lower chambers were filled with serum-free medium, and 200,000 cells were added to the upper chamber. After the cells were allowed to settle for 1 h, 1 μg/ml CCL25 (PeproTech) was added to the lower chamber and cell migration was measured by counting the cells in the lower chamber 3 h after the addition of CCL25. Specific cell migration was calculated by subtracting the number of cells migrating in the absence of chemokine from the number of cells migrating in the presence of chemokine and then calculating the proportion of cells in the lower chamber as a percentage of the number of input cells.

Adoptive T-cell transfer.For autologous RM-1 and RM-2 CCR9-transduced T-cell infusion cultures, the cells to be infused were labeled with 5 μM CTV (Invitrogen) according to the manufacturer’s procedure. For the animal RM-2 infusion product, the parallel unmodified CD8 T-cell culture was labeled with 5 μM CTFR (Invitrogen) and mixed with the CVT-labeled transduced culture to produce the RM-2 infusion product. The respective infusion products were washed in PBS, resuspended in 50 ml saline solution with 2% (vol/vol) autologous serum, and infused into the saphenous vein (flow rate of 2 ml/min) as previously described (11, 51, 52). To monitor the transfer of cells, preinfusion and 15-min postinfusion samples were collected from the cephalic vein.

Confocal microscopic tissue analysis.Tissue was embedded in Optimal Cutting Temperature compound (OCT; Tissue Tek), thin sectioned, and placed in 4% (vol/vol) paraformaldehyde-water for 15 min at 4°C, followed by a brief submersion in ethanol and air drying. Fixed sections were incubated in a 1:500 dilution of anti-CCR9 (38097; Dako) in Tris-buffered saline (Boston Biosciences) with 0.05% (vol/vol) Tween 20 and 0.25% (wt/vol) casein for 1 h at room temperature and then incubated in a 1:500 dilution of Alexa 594-conjugated donkey anti-rabbit secondary antibody for 15 min at room temperature. Slides were rinsed and mounted with number 1.5 Gold Seal cover glasses (Electron Microscopy Services) using Prolong Gold mounting medium (Invitrogen). Regions of interest were acquired using an Olympus FluoView FV10i system at ×100 for overview and ×600 for close-up magnification.

ACKNOWLEDGMENTS

Within the AIDS and Cancer Virus Program of the Frederick National Laboratory, we thank the members of the Nonhuman Primate Research Support Core, Vicky Coalter, Don Johnson, Jacob Kiser, George Muthua, Adam Wiles, and Rodney Wiles, for expert sample collection, processing, and technical assistance; Simona Florea, Leslie Johnston, and David Morcock of the Tissue Analysis Core for technical tissue analysis; Cathi Pyle and James Thomas for absolute count flow cytometry; and the Laboratory Animal Science Program, Building 14D South, animal technicians, caretakers, and management staff for their expert technical support and excellent animal care. We also acknowledge the National Institutes of Health (NIH) Nonhuman Primate Reagent resource for providing the α4β7 antibody, number PR-1427.

This project has been funded in whole or in part with federal funds from the National Cancer Institute, National Institutes of Health, under contract no. HHSN261200800001E. The content of this publication does not necessarily reflect the views or policies of the Department of Health and Human Services, and mention of trade names, commercial products, or organizations does not imply endorsement by the U.S. Government.

FOOTNOTES

    • Received 28 May 2019.
    • Accepted 16 August 2019.
    • Accepted manuscript posted online 21 August 2019.

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KEYWORDS

adoptive cell therapy
adoptive cell transfer
CCL25
CCR9
immunology
infused cell persistence
SIV
small intestine
T cell homing
T cell immunotherapeutics

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