Identification and Characterization of pDC-Like Cells in Normal Mouse Skin and Melanomas Treated with Imiquimod

Francesca Palamara, Simone Meindl, Martin Holcmann, Petra Lührs, Georg Stingl and Maria Sibilia
J Immunol September 1, 2004, 173 (5) 3051-3061; DOI: https://doi.org/10.4049/jimmunol.173.5.3051

Abstract

Among the different subsets of dendritic cells (DC) described in humans and mice, epidermal Langerhans cells and dermal DCs represent the only DC populations resident in normal skin. In this study we describe a population of CD4+CD3− plasmacytoid DC (pDC)-like cells that accumulate in the dermis and spleens of mice topically treated with imiquimod, a low m.w. immune response modifier with potent antiviral and antitumor activities. These CD4+CD3− cells coexpress GR-1, B220, MHC class II, and, to a lesser extent, CD11c and display the phenotypic features of pDCs described in lymphoid organs. The accumulation of pDC-like cells after imiquimod treatment was detected not only in normal skin, but also in intradermally induced melanomas. Imiquimod treatment leads either to complete regression or to a significant reduction of the tumors. The number of pDCs correlates well with the clinical response of the tumors to the drug, suggesting that the antitumor effects of imiquimod could be mediated at least in part by the recruitment of pDC-like cells to the skin. Therefore, strategies aimed at activating and directing these cells into neoplastic tissues may be a promising and novel approach for the immunotherapy of various types of cancer.

Dendritic cells (DC)4 constitute a heterogeneous population of APCs found in virtually every tissue and organ. They link innate and adaptive immunity by inducing appropriate immune responses upon the recognition of invading pathogens, thus acting as “nature’s adjuvant” (1, 2). In peripheral tissues DCs continuously monitor the antigenic environment, and any encounter with microbial products or damaged tissue induces the activation and the migration of DCs to the draining lymph nodes (LN). In LN, DCs present Ags to T lymphocytes and then efficiently trigger immune responses through the selective binding between TCRs and the foreign peptides complexed on the DC surface to MHC (1, 2). Many subsets of DC have been described in both humans and mice (1, 3, 4, 5). Among these, epidermal Langerhans cells (LC) and dermal DC have been shown to be the only resident DC populations in normal skin. Recently, another subset of DC, termed plasmacytoid DC (pDC), has been described in human skin, occurring in cutaneous inflammatory diseases such as psoriasis vulgaris, contact dermatitis, and lupus erythematosus and being almost absent in normal skin (6, 7).

Originally pDCs were identified in humans as a population of medium-sized cells readily identifiable in tissue sections on the basis of their plasma cell-like morphology (8). They were initially termed T-associated plasma cells or plasmacytoid T cells, because they do not express the typical markers of plasma cells and are localized in the T cells zone of lymphoid tissues. After this discovery, a series of studies has phenotypically characterized these cells in humans as Lin− DC (9, 10, 11, 12). It has been shown that human pDCs produce large amounts of type I IFNs (IFN-α and IFN-β) and thus form a key component of the antiviral defense strategy of the immune system. They seem to be identical with the natural type I IFN-producing cells (10, 11, 13), which are capable of synthesizing extremely high amounts of type I IFN upon viral infection (14), and to develop characteristic features of DCs upon maturation (10, 15). Human pDCs can be identified by the coexpression of CD123 (IL-3R) and CD45RA, BDCA-2, CD68, CXCR3, and CD62L (L-selectin). They also express CD4 and MHC class II (MHC-II), but not CD11c or CD45RO (5, 10).

In the last few years several groups simultaneously identified the murine equivalent of human pDCs as CD11c+B220+GR-1+MHC-II+ cells of plasmacytoid morphology, producing large amounts of type I IFN and therefore also termed IFN-producing cells (16, 17, 18, 19, 20). Mouse pDCs appear to be negative for markers such as CD11b, CD19, CD40, CD80, CD86, CD79b, and F4/80 and positive at various degrees for CD45RA, CD45RB, CD123, and CD62L (12, 21). Moreover, different groups found pDCs to be CD4+CD8α−, CD4−CD8α+, or CD4+CD8α+ depending on the mouse strain, localization, or activation state (5, 12, 17, 18, 19, 20, 21, 22, 23, 24, 25). These cells have been detected in peripheral blood, bone marrow, LN, and spleen, but not in nonlymphoid tissues.

Recently, a family of type I transmembrane receptors termed TLRs has been described that seems to be involved in the recognition of pathogen-associated molecular patterns, such as LPS, peptidoglycans, bacterial DNA containing unmethylated CpG motifs, and dsRNA (26, 27). The identification of TLRs revealed how innate immunity is closely linked to acquired immunity (26, 27). Among the 10 family members of TLRs (TLR1 to TLR10) identified to date, mouse pDCs seem to express only TLR7 and TLR9 and to respond to TLR7 and TLR9 ligands by producing large amounts of IFN-α (26, 28, 29). Although TLR9 seems to be essential for the recognition of bacterial DNA containing unmethylated CpG motifs, the natural ligand for TLR7 was still unknown until very recently, when two groups reported that ssRNAs represent the physiological ligands for murine TLR7 and human TLR7 and -8 (30, 31).

TLR7 is also selectively recognized by small synthetic immune response modifiers, including imiquimod, R-848, loxirabine, and bropirimine (29, 32). Imiquimod has potent antiviral and antitumor activities (32, 33, 34, 35) and can induce the synthesis of IFN-α and other cytokines (TNF-α, IL-1, IL-6, IL-8, IL-12, GM-CSF, etc.) in a variety of cell types (36, 37, 38). Recently, imiquimod became available for the treatment of human papillomavirus infections, particularly for genital and perianal warts. It has also been shown to be efficient in the treatment of infectious diseases such as genital herpes virus and CMV infections in guinea pigs (39, 40). Moreover, imiquimod can act as a potent antitumor drug in a number of murine tumor models, including MC-26 colon carcinoma, Lewis lung carcinoma, and FCB bladder tumor (41, 42). Topical application of imiquimod to mouse skin induces a strong inflammatory response, accompanied by alterations in LC morphology, enhancement of LC migration to the draining LN, and up-regulation of different cytokines in the skin (IL-6, IL-10, IFN-α, IFN-γ, and TNF-α) (43).

In this study we aimed at obtaining more detailed information about the quality and quantity of the leukocytic infiltrate induced by imiquimod. We were also interested to determine whether imiquimod, similarly to the human situation, can induce the regression of superficial melanocytic neoplasms of the skin, and if so, whether this phenomenon can be correlated with a particular phenotypic profile of leukocytes invading and surrounding the tumor.

Materials and Methods

Mice, cells, and reagents

The compound imiquimod (1-(2-methylpropyl)-1H-imidazo[4,5]quinoline-4-amine) used in this study is a proprietary molecule of 3M Pharmaceuticals (St. Paul, MN), and the structure of this molecule has been described previously (44). Female C57BL/6 (Haarlan-Winkelmann, Borchen, Germany) mice between 8 and 12 wk of age were treated with ∼10–12 μl of a 5% imiquimod cream formulation (Aldara). The cream was either applied at the dorsum of one ear or to the shaved back skin once daily for 8 days. As a negative control, the contralateral ears or the back skin were treated with vehicle.

Melanoma cells M3 (derived from Cloudman S91 melanoma; H-2d) were cultured in Ham’s F-10 standard medium as previously described (45). M3 cells (6 × 105/mouse) were then injected intradermally in previously shaved back skin of anesthetized DBA/2 female mice (Haarlan-Winkelmann). Tumor growth was monitored every 2–3 days. Melanomas that had reached a certain volume were treated topically with either imiquimod cream (5%) or vehicle every 2–3 days over a period of 30 days (total of 12 applications), and the sizes of the tumors were measured every 3–4 days by tissue calipers. At the end of each treatment, mice were killed by cervical dislocation, and the tissues were processed further.

Antibodies

The following purified mAbs were purchased from BD Pharmingen (San Diego, CA): anti-CD3ε, PE-conjugated anti-CD4, R-PE-conjugated anti-CD8α, R-PE-conjugated anti-CD11c, FITC-conjugated anti-CD45, anti-CD19, FITC- and R-PE-conjugated anti-I-A/I-E, and FITC-conjugated anti-Ly-6G (Gr-1) and -Ly-6C. The respective isotype controls were also obtained from BD Pharmingen. Anti-F4/80 was purchased from Serotec (Kidlington, U.K.). Second-step, biotin-conjugated, goat α-hamster IgG and rabbit α-rat IgG Abs (Vector Laboratories, Burlingame, CA) were used at the recommended dilutions. Secondary fluorescent Abs were purchased from Molecular Probes (Eugene, OR): goat α-rat IgG Alexa-Fluor 488 (green), goat α-rat IgG Alexa-Fluor 546 (red), and goat α-hamster IgG Alexa-Fluor 488 (green). Streptavidin-Cy5 was purchased from Jackson ImmunoResearch Laboratories (West Grove, PA).

Immunolabeling of Langerhans cells

Skin samples were collected between days 0 and 20 after imiquimod and vehicle treatment, and anti-MHC-II in situ immunolabeling was performed as previously described (46). Positively stained cells were counted by examining at least 10 randomly selected fields in each group, and their morphology was analyzed under a fluorescence microscope. Results are expressed as positively stained cells per square millimeter (mean ± SEM). The experiment was repeated three times.

Immunohistology

Mice were killed, and tissues were either snap-frozen in liquid nitrogen for cryosections or fixed in 4% PBS-buffered formaldehyde and embedded in paraffin. Five-micron sections were cut and stained either with H&E (Sigma-Aldrich, St. Louis, MO) according to standard procedures or were processed further for immunohistochemical or immunofluorescence staining. Immunohistochemical stainings were performed using the avidin-biotin-peroxidase staining kit (Vector Laboratories) according to the manufacturer’s recommendations, and immunofluorescence stainings of frozen sections were performed as described previously (7). In all staining experiments, substitution of the primary Abs with isotype-matched IgG and omission of the primary Ab served as negative controls. Positively stained cells were counted by examining at least 10 random fields in each group and were expressed as number of cells per field (mean ± SEM; magnification, ×200). The statistical significance of the data was determined by applying the two-tailed Student’s t test. The difference was considered statistically significant at p < 0.05.

Preparation of dermal and spleen cell suspensions and FACS analysis

Single-cell suspensions from skin and/or spleen of imiquimod- and vehicle-treated mice were prepared for flow cytometric analysis (FACS) as follows. Mice were killed on day 8 after treatment, and skin/ear samples were collected and cut into pieces. Trypsin/PBS 0.8% (30 min, 37°C) was used to separate the epidermis from the dermis. Samples of dermis were collected and incubated for 45 min at 37°C in DNase medium (200 mg/ml DNase I; Sigma-Aldrich) and then passed through a 70-μm pore size nylon mesh. Spleen samples were minced through a 70-μm pore size nylon mesh, and the cells were collected and incubated for 15 min at 4°C with 2% PBS/BSA supplemented with 10% mouse serum and 5% goat serum to block FcγIII/IIR (CD16/CD32). Subsequently, cells were stained for the indicated markers and analyzed by flow cytometry (FACSCalibur; BD Biosciences, Mountain View, CA); matched isotype mAb served as controls. Dead cells positive for 7-aminoactinomycin were excluded. Two hundred thousand events per sample were acquired, and 5% of them were displayed (10,000). All experiments were repeated five times.

RT-PCR analysis of enriched splenic DC populations

Isolated spleens of vehicle- and imiquimod-treated mice were injected with HBSS buffer containing collagenase D (400 U/ml), cut into small pieces, incubated for 20 min at room temperature, and subsequently passed through a 70-μm pore size nylon mesh. The collected cells were centrifuged, resuspended in 2 ml Lymphoprep (Tecnoclone, Vienna, Austria), overlaid with 2 ml of HBSS, and centrifuged for 15 min at 1700 × g. The low density fraction enriched for DCs was collected, and RNA was extracted with the RNeasy Minikit (Qiagen, Valencia, CA). After DNase I digestion (Invitrogen Life Technologies, Carlsbad, CA), RT was performed from 1.3 μg of RNA of each sample using Superscript RT-II (Invitrogen Life Technologies). PCR was performed on the cDNAs with the following primers: IFN-α: forward, TGTCTGATGCAGCAGGTGG; and reverse, AAGACAGGGCTCTCCAGAC (47); TLR7: forward, TGACTCTCTTCTCCTCCA; and reverse, GCTTCCAGGTCTAATCTG (48); TNF-α: forward, AGTGGTGCCAGCCGATGGGTTGT; and reverse, GCTGAGTTGGTCCCCCTTCTCCAG; IL-2: forward, ATGTACAGCATGCAGCTCGCATC; and reverse, GGCTTGTTGAGATGATGCTTTGACA; IL-4: forward, TCGGCATTTTGAACGAGGTC; and reverse, GAAAAGCCCGAAAGAGTCTC (49); IL-5: forward, TCACCGAGCTCTGTTGACAA; and reverse, CCACACTTCTCTTTTTGGCG (49); IL-6: forward, ATGAAGTTCCTCTCTGCAAGAG; and reverse, CCAGTTTGGTAGCATCCATC; IL-23: forward, ATAATGTGCCCCGTATCC; and reverse, ACAAACGAAACAAGAACAGC; and hypoxanthine phosphoribosyltransferase: forward TTGCTCGAGATGTGATGAAGGA; and reverse, AAAGTTGAGAGATCATCTCCACCAA.

Results

Imiquimod-induced migration of LC is reversible

Topical administration of imiquimod on normal mouse skin induces emigration of LCs from the epidermis (43). To confirm these findings and to investigate whether these effects are reversible after termination of treatment, anti-MHC-II in situ immunolabeling was performed on epidermal sheets of imiquimod- and vehicle-treated mouse ears. Whereas vehicle treatment had no effect on the number of LCs present in the epidermis, the number of LCs per square millimeter started to decrease after 3 days of imiquimod treatment, falling to ∼40% of the control value by day 8 when the treatment was stopped (Fig. 1⇓A). On day 10 (2 days after the last treatment), the number of LCs in the epidermis was still greatly reduced, but they reached their original number by day 20 (Fig. 1⇓A). The decrease in the number of epidermal LCs in imiquimod-treated ears was associated with a change in their morphology (Fig. 1⇓, B and C). By day 8, the LCs remaining in the epidermis after imiquimod treatment appeared larger and more dendritic and stained more strongly for MHC-II Ag than LCs present in vehicle-treated skin (Fig. 1⇓C and data not shown). These morphological changes persisted throughout the treatment until day 10 and are indicative of an activated state. On day 20, the morphology of LCs of imiquimod-treated skin again appeared comparable to that of vehicle-treated controls (Fig. 1⇓D and data not shown). These results show that LC migration induced by imiquimod is reversible after termination of treatment.

FIGURE 1.
FIGURE 1.

Effect of imiquimod on LC density and morphology. A, Number of MHC-II+ LC present in epidermal sheets of mouse ears treated once daily with 10–12 μl of Aldara cream (5%) or vehicle for 8 days. The arrow indicates the end of imiquimod/vehicle treatment. The data represent the mean ± SEM of the number of MHC-II+ cells counted in 10 fields of two independent samples. Immunofluorescence staining of epidermal sheets on day 0 (B), day 8 (C), and day 20 (D) after beginning of imiquimod treatment is shown. Magnification, ×200. ∗, p < 0.05.

Application of imiquimod leads to skin inflammation

To better investigate the effects of imiquimod on the cutaneous immune system, histological and immunohistochemical examinations were performed on skin on day 8 after treatment. Macroscopic examination and measurement of skin thickness by a mechanical micrometer showed inflammation, edema, and swelling starting around day 3 after imiquimod treatment (data not shown). These changes were not detectable in vehicle-treated controls in which the skin appeared normal and comparable to that in nontreated controls (data not shown). Application of imiquimod induced spongiosis, acanthosis, and papillomatosis as well as pronounced inflammatory changes in the dermis that were never observed in vehicle-treated skin (Fig. 2⇓, A and B). Immunohistochemical staining of imiquimod-treated skin showed a massive increase in CD45+ leukocytes, which consisted, in decreasing order, of MHC-II+, CD4+, CD11c+, GR-1+, Mac-1+, F4/80+, and CD3+ cells (Fig. 2⇓C). Only a few CD8α+ cells were present, and their number was not significantly different from that in vehicle-treated skin (Fig. 2⇓C). Staining for CD19 showed that both vehicle- and imiquimod-treated skin were almost devoid of B cells (Fig. 2⇓C). Interestingly, the number of CD4+ cells regularly exceeded that of CD3+ cells, suggesting that some of these CD4+ cells are not T cells (Fig. 2⇓C). These results show that topical application of imiquimod induces skin inflammation with increased numbers of inflammatory cells in the dermis.

FIGURE 2.
FIGURE 2.

Topical imiquimod treatment leads to skin inflammation. A and B, Imiquimod- and vehicle-treated skin samples were collected on day 8 after beginning of treatment (once daily for 8 days), and cryosections were stained with H&E. B, Pronounced inflammatory changes such as acanthosis and papillomatosis (marked on the right of B) and leukocytic infiltrates (boxed area in B) were present in the dermis of imiquimod-treated skin. Similar changes were not seen in control skin (A). The dotted lines in A and B delineate the dermal-epidermal junction. Magnification, ×200. C, Immunohistochemical stainings performed on imiquimod- and vehicle-treated back skin sections on day 8 after beginning of treatment (once daily for 8 days) showing the numbers of stained cells present in the respective skin samples. The data represent the mean ± SEM of the number of positive cells counted in seven fields of three independent samples. ∗, p < 0.05.

Characterization of the inflammatory infiltrate of imiquimod-treated skin

It has been shown that imiquimod acts via binding to TLR7 and that in mice this receptor appears to be selectively expressed on pDCs (29, 50). Moreover, mouse pDCs seem to express CD4 on their surface (5, 12, 24). Therefore, we next characterized the inflammatory infiltrate of imiquimod-treated skin, searching particularly for the presence of phenotypic markers of pDCs. Immunofluorescence double labeling revealed the presence of a population of CD4+CD3− cells in both treated and nontreated skin (Fig. 3⇓A). In imiquimod-treated skin, these CD4+CD3− cells appeared larger, exhibited a round shape, and were greatly increased in number (Fig. 3⇓A and data not shown). Moreover, these CD4+ cells were also positive for markers such as CD11c (Fig. 3⇓, A–C), GR-1 (Fig. 3⇓, A, D, and E), and MHC-II (Fig. 3⇓A). All these double-positive cells as well as cells coexpressing CD11c+GR-1+, MHC-II+CD11c+, and MHC-II+GR-1+ were significantly increased in number in imiquimod-treated skin (Fig. 3⇓A). These results indicate that CD4+CD3− cells present in imiquimod-treated skin display the features of pDC expressing on their surface markers such as MHC-II, GR-1, and, to a lesser extent, CD11c.

FIGURE 3.
FIGURE 3.

Characterization of the inflammatory infiltrate in the skin of mice after topical application of imiquimod. Double-immunofluorescence stainings with the indicated Abs were performed to phenotypically analyze the leukocytic infiltrate. A, The number of single- and double-positive cells for the respective markers is displayed. The data represent the mean ± SEM of the number of single- and double-positive cells counted in 10 fields of three independent samples. ∗, p < 0.05. B–E, A pronounced increase in the number of dermal CD4+CD11c+ (C) and CD4+GR-1+ (E) cells was observed in imiquimod-treated skin compared with control skin (B and D). Magnification, ×200. g, green, FITC-labeled Abs; r, red, R-PE-labeled Abs.

FACS analysis of dermal cell suspensions from imiquimod- and vehicle-treated skin also revealed a significant increase in the number of MHC-II+GR-1+ and CD45RB+GR-1+ cells after imiquimod treatment (Fig. 4⇓). We also found an increase in the number of B220+MHC-II+ and B220+GR-1+ cells in imiquimod-treated skin (Fig. 4⇓). These B220+ cells are probably not B lymphocytes, but, rather, pDCs, because by immunohistochemical staining the number of CD19+ B cells was very low after imiquimod treatment (Fig. 2⇑C). Given that B220 is expressed on murine pDCs (20), these findings suggest that pDCs are present and increased in number in imiquimod-treated skin.

FIGURE 4.
FIGURE 4.

Analysis of the dermal inflammatory infiltrate of imiquimod-treated skin. Flow cytometric (FACS) analysis of dermal single-cell suspensions showing an increase in the number of B220+MHC-II+, B220+GR-1+, MHC-II+GR-1+, and CD45RB+GR-1+ cells in imiquimod-treated skin (right) compared with vehicle-treated controls (left).

Splenomegaly and increase in splenic pDC-like cells in imiquimod-treated mice

When mice that had been treated daily with imiquimod for 8 days were killed, we regularly observed a pronounced splenomegaly (Fig. 5⇓A). FACS analysis of cell suspensions prepared from such spleens revealed that although the number of T and B cells was decreased, the number of DCs, NK cells, granulocytes, neutrophils, and macrophages was significantly increased (Fig. 5⇓B). Among the DC population there was a significant increase in the number of pDC-like cells in spleens of imiquimod-treated mice. These pDC-like cells were triple positive for CD11c/GR-1/B220, B220/GR-1/MHC-II, and B220/GR-1/CD4 as well as CD11c/MHC-II/B220, CD4/CD11c/B220, and CD11c/F4/80/B220 (Fig. 5⇓C and data not shown). To investigate whether cytokines known to be induced by imiquimod were expressed at increased levels, RNA was extracted from splenic cell populations enriched for DCs and analyzed by semiquantitative RT-PCR. Interestingly, TLR7 and IFN-α, which are both known to be expressed by mouse pDCs, were already expressed in resident spleen cells, and their expression was slightly up-regulated after imiquimod treatment (Fig. 5⇓D). In addition, several cytokines, such as TNF-α, IL-2, IL-4, IL-5, and IL-6, were expressed at increased levels in spleen cells after imiquimod treatment (Fig. 5⇓D). Taken together, these results show that topically applied imiquimod can have systemic effects, leading to splenomegaly with an overproportional increase in pDC in this organ.

FIGURE 5.
FIGURE 5.

Analysis of spleen cells of imiquimod-treated mice. A, Macroscopic view of the spleen of vehicle-treated (left) and imiquimod-treated (right) mice showing splenomegaly after topical imiquimod application. B, The percentages of the different cell populations detected in the spleen after FACS analysis for the indicated markers are shown (granul., granulocytes; neutr., neutrophils). C, FACS analysis (triple stains) of splenic single-cell suspensions, demonstrating an increase in the number of CD11c+GR-1+B220+, B220+GR-1+MHC-II+, and B220+GR-1+CD4+ cells in imiquimod-treated mice compared with vehicle controls. The percentages of total cells present within the boxed regions are indicated. In the first row, blue and red dots and numbers refer to CD11c− and CD11c+ cells, respectively. D, RT-PCR analysis showing the expression of TLR7 and several cytokines in DC-enriched spleen cell populations. V, vehicle; I, imiquimod; +/− RT, with and without reverse transcriptase.

Effect of imiquimod on melanoma growth

To investigate the antitumor properties of imiquimod in vivo, we used a melanoma model in which DBA/2 mice were injected intradermally with 6 × 105 M3 melanoma cells. After a period of 16 ± 7 days, 57% of the injected mice had developed melanomas (Fig. 6⇓A). The skin overlying and surrounding the tumors was treated with either vehicle or imiquimod once a day every second day for 30 days. Vehicle treatment did not affect tumor growth, and tumors had considerably increased in size after 30 days (Fig. 6⇓, A, B, and E–G). Overall, in 26% of imiquimod-treated mice there was complete tumor regression (Fig. 6⇓, A and H-J), in 51% of the mice the tumor size remained essentially unchanged throughout the entire treatment period, and in 23% of mice there was poor response to the drug, and tumors continued to grow (Fig. 6⇓A). The clinical response of the tumors to imiquimod very much depended on the size of the melanomas at the beginning of the treatment. When the tumor size was small (8–180 mm3), the response to imiquimod was much better, and there was complete tumor regression in ∼38% of the mice, stable disease in 58%, and tumor progression in only 4% of the mice (Fig. 6⇓, A and C). In contrast, when tumor size at the beginning of the experiment ranged from 180-1200 mm3, only 7% of the mice showed complete regression, whereas 40% of the mice showed stable disease, and 53% showed progressive disease (Fig. 6⇓, A and D). However, even these progressor tumors never reached the size of those treated with vehicle, suggesting that imiquimod still had a tumoricidal effect in this situation (Fig. 6⇓D). These results show that imiquimod treatment inhibits tumor growth and that this inhibition is more effective when the size of the melanomas is small at the beginning of treatment.

FIGURE 6.
FIGURE 6.

Effect of imiquimod on melanoma growth. A, DBA/2 mice were injected intradermally with 6 × 105 M3 melanoma cells. Of 115 injected mice, 65 (57%) developed melanomas, which were either treated with vehicle (n = 26) or imiquimod (n = 39) every 2–3 days for 4 wk (total of 12 applications). The total number of mice and the respective percentages showing the various clinical responses after imiquimod and vehicle treatments are indicated. Mice treated with imiquimod (n = 39) were further subdivided according to the tumor volume present at the beginning of treatment (small and big tumors), and the number of mice and respective percentages showing clinical responses are displayed. B, Tumor growth curve, representing the relative tumor volumes of all imiquimod- and vehicle-treated mice (small and big tumors together). C and D, Tumor growth curves represented as relative tumor volumes of imiquimod- and vehicle-treated mice grouped according to small (between 8 and 180 mm3; C) and big (between 180 and 1200 mm3; D) tumor sizes. Results are representative of three independent experiments. ∗, p < 0.05. E–G, Macroscopic appearance of tumors treated with vehicle at the indicated time points. H–J, A mouse showing tumor shrinkage starting around day 12 after imiquimod treatment (I), with complete regression by day 15 (J). V, vehicle; I, imiquimod.

To investigate the mechanism of growth inhibition, the number of proliferating and apoptotic cells was measured by immunohistochemical stainings for Ki67 and caspase-3, respectively. Ki67 staining of tumor tissue revealed an ∼50% reduction in the number of proliferating cells in tumors treated with imiquimod (94 ± 8) compared with tumors treated with vehicle (196 ± 60). In imiquimod-treated tumors, almost twice as many apoptotic cells were detected (34 ± 17) relative to vehicle-treated controls (17 ± 11), indicating that imiquimod leads to reduced proliferation and increased apoptosis of tumor cells. Histologically, melanomas of vehicle-treated mice consisted mostly of large tumor cells with lots of leukocytes surrounding them, but only a few of them infiltrating the tumor tissue (Fig. 7⇓, A and B). In contrast, a massive leukocytic infiltrate could be detected in imiquimod-treated tumors (Fig. 7⇓, C–F). In fact, imiquimod-treated tumors, which were classified as nonresponders, also contained high numbers of leukocytes surrounding the tumor cell clusters compared with vehicle-treated controls (Fig. 7⇓, C and D). Even when tumors had completely disappeared in response to imiquimod treatment, leukocytic infiltrates including mast cells and activated fibroblasts could be detected in the dermis (Fig. 7⇓, E and F). These results show that imiquimod treatment leads to the appearance of inflammatory cells that invade tumor tissue.

FIGURE 7.
FIGURE 7.

Analysis of imiquimod- and vehicle-treated melanomas. A, Histological examination of a vehicle-treated tumor on day 15 showing large melanoma cells infiltrating the entire dermis and the s.c. fat tissue. Leukocytes (arrows) are present in the region of the dermis surrounding the tumor. B, Higher magnification of the area boxed in A, showing that no inflammatory cells were detected in the tumor mass. The arrows point to large melanoma cells forming a compact tumor mass. C and D, Imiquimod-treated tumor (nonresponder) on day 23 of treatment, displaying large numbers of leukocytes (arrowheads in D) infiltrating the tumor tissue. Melanoma cells (arrows in D) appear smaller and fewer in number compared with controls (B). E and F, Imiquimod-treated tumor (responder) on day 15 of treatment, when no tumor cells were detected in dermis or s.c. fat tissue. Some leukocytes (arrowheads in F) and activated fibroblasts (evidenced by strong pink eosin staining in E and F) were present in the dermis. Magnification: A, C, and E, ×100; B, D, and F, ×400.

Characterization of the inflammatory infiltrate in imiquimod-treated melanomas

Computer-assisted morphometric analysis revealed that compared with the vehicle-treated control, the total skin area covered by CD45+ leukocytes was significantly increased after imiquimod treatment (data not shown). This increase was more pronounced in regressor tumors and in mice with stable disease (data not shown). To determine whether pDCs-like cells were present in the tumor inflammatory infiltrate, sections from imiquimod- and vehicle-treated melanomas were analyzed and stained for the main markers of mouse pDCs.

We found that vehicle-treated tumors displayed a sizable number of CD4+CD3+, but few CD4+CD3− cells, indicating that the majority of CD4+ cells are T cells (Fig. 8⇓A). In contrast, in imiquimod-treated tumors, higher numbers of CD4+CD3− cells were detected, confirming the findings already made in normal mouse skin treated with imiquimod (Fig. 8⇓, B and I). Not many CD4+MHC-II+ double-positive cells were seen in vehicle-treated melanomas, whereas they could be detected at higher numbers after imiquimod application (Fig. 8⇓, C and D). Only a few GR-1+ cells were found in control tumors, and not many of them appeared to coexpress markers such as CD4, MHC-II, and CD11c (Fig. 8⇓, E, G, and I). In contrast, many GR-1+CD4+, GR-1+MHC-II+, as well as GR-1+CD11c+ double-positive cells were found in imiquimod-treated tumors (Fig. 8⇓, F, H, and I), whereas MHC-II+CD11c+ cells were present in both imiquimod- and vehicle-treated tumors (data not shown). Interestingly, the area covered by CD8+ T cells was always much higher in vehicle-treated tumors even when compared with nonresponder, imiquimod-treated melanomas (data not shown). These results indicate that only imiquimod and not vehicle application is followed by an increased appearance of pDC-like cells in and around the tumors. The number of these cells correlates well with the clinical response of the tumors to the drug being higher in responders and in mice with stable disease. These results suggest that the antitumor effects of imiquimod could be mediated in part by the recruitment of pDC-like cells to the skin.

FIGURE 8.
FIGURE 8.

Characterization of the inflammatory infiltrate in imiquimod- and vehicle-treated melanomas. Cryosections from imiquimod- and vehicle-treated melanomas on day 19 of treatment (total of eight treatments) were stained with the indicated Abs. A and B, A high number of CD4+CD3− cells (red) were detectable in imiquimod-treated tumors (B), whereas in controls almost all the CD4+ cells also stained for CD3 (yellow, A). C and D, CD4+MHC-II+ cells (yellow) were only found in imiquimod-treated skin (D), not in vehicle-treated controls (C). E–H, Few GR-1+ cells (green), which were negative for CD4 (E) and MHC-II (G), were detected in control tumors, whereas many GR-1+CD4+ (F) and GR-1+MHC-II+ (H) double-positive cells (yellow) were found in imiquimod-treated melanomas. Magnification, ×400. g, green, FITC-labeled Abs; r,: red, R-PE-labeled Abs. I, The percentage of double-positive cells of the respective total single-positive cells is displayed for the indicated markers. The numbers of single- and double-positive cells were determined by counting 10 fields of two independent samples.

Discussion

The immune response modifier imiquimod has been shown to exert antiviral and antitumor activities in both animal models and man (32, 40, 51, 52, 53). Imiquimod is a potent cytokine inducer (33, 37, 38, 54), and this activity is at least in part responsible for its clinical effects. It has recently been shown that topical application of imiquimod leads to LC activation and their migration to draining LN (43). In our study we observe similar effects, and we further demonstrate that the migration of LC in response to imiquimod is dose dependent and reversible after termination of treatment. The degree of LC migration induced by imiquimod is comparable to what was observed after LPS treatment. In the case of LPS it seems that the migratory effect on LCs is mediated indirectly via the release of cytokines such as TNF-α (55). It is unclear whether imiquimod-induced migration is due to a direct effect of imiquimod on LCs or to the action of the drug on a different target cell, which releases cytokines that, in turn, lead to LC activation and migration. Because it has been shown that imiquimod as well as its more potent analog R-848 are selectively recognized by TLR7, mainly expressed on pDCs in mice (29), we investigated whether the mechanism of action of imiquimod may involve the recruitment of different populations of leukocytes to the skin, looking particularly for the presence of phenotypic markers of murine pDCs.

Interestingly, a population of CD4+CD3− cells, displaying characteristic features of mouse pDCs observed in lymphoid organs (3, 5, 12, 21, 22, 23, 25), was present and significantly increased in number in mouse skin after imiquimod treatment. Most of these CD4+ cells coexpressed markers such as GR-1, MHC-II, and, to a lesser extent, CD11c, supporting our hypothesis that these cells might represent murine pDCs. Indeed, FACS analysis also revealed the presence of high numbers of B220+GR-1+, B220+MHC-II+, MHC-II+GR-1+, and CD45RB+GR-1+ cells in the dermis of treated skin. It is likely that the B220+MHC-II+ cell population observed in treated skin consists mostly of pDCs rather than B cells because we already observed that CD19+ B lymphocytes are only present at very low numbers in the skin and are not affected by imiquimod treatment. Simultaneous coexpression of B220, MHC-II, and CD45RB has never been reported on granulocytes (GR-1+), suggesting that these cells appearing in the skin after imiquimod treatment are indeed mouse pDCs, which have been shown to be positive for markers such as CD4, GR-1, B220, MHC-II, and CD11c (12, 18, 19, 20, 21, 23). According to these findings, we propose these cells to represent a population of pDCs and we term them cutaneous pDC-like cells.

Costainings with CD4 could not be performed on skin by FACS analysis, because optimal dermal cell suspensions could only be obtained after trypsin digestion, and it is known that the CD4 Ag is trypsin sensitive. However, FACS stainings with CD4 Ag could be performed in spleens of imiquimod-treated mice. These spleens appeared bigger after topical application of imiquimod, most likely due to systemic effects induced by the drug, because the permeability of imiquimod through the skin seems to be 40 times higher in mice than in men (H. B. Slade, unpublished observation). The number of macrophages, NK cells, and DCs was markedly increased in spleens of mice topically treated with imiquimod. Among the DC population, the number of pDC-like cells triple positive for CD4/GR-1/B220 and CD4/CD11c/B220 was significantly increased by imiquimod. Moreover, we observed that the expression of TLR7 and several cytokines known to be induced by imiquimod was increased after imiquimod treatment. Taken together, these findings demonstrate that imiquimod treatment leads to an increase, in spleen and skin, of a population of cells displaying all the characteristic features of mouse pDCs expressing on their surface markers such as CD4, CD11c, MHC-II, GR-1, and B220.

At the moment it is still unclear whether the appearance of pDC-like cells in skin and that in spleen are independent or linked events, and several possibilities can be entertained to explain their origin. They could be recruited into the skin as a primary effect of imiquimod and then migrate into the spleen. Alternatively, they could be increased in number in the spleen as a consequence of a systemic effect of the molecule binding to TLR7 expressed on resident splenic pDCs and then migrate into the skin, or finally, the effects observed in spleen and skin could be independent of each other and due to topical and systemic effects, respectively. The first and the third possibilities would imply the existence, until now never shown, of some resident or peripheral blood-derived precursors in the skin expressing TLR7 and able to proliferate in response to the drug. The second possibility would be easily explained by the evidence that pDC expressing TLR7 are usually present in the spleen under normal conditions (17). After imiquimod treatment, pDC precursors resident in the spleen could be induced to proliferate and migrate into peripheral organs. However, it can also be assumed that a combination of these possibilities leads to the observed phenotypes. The question that remains to be resolved is whether imiquimod, after binding to TLR7 on splenic and/or cutaneous pDCs themselves, would induce their influx from the blood. Alternatively, imiquimod could bind to either TLR7 or an as yet unidentified receptor structure on cells other than pDCs and indirectly trigger the attraction of pDCs into the tissues of interest.

We also investigated whether imiquimod, similarly to the human situation (51, 52, 53), can induce the regression of superficial melanocytic neoplasms of the skin and, if so, whether this phenomenon can be correlated with a particular phenotypic profile of leukocytes invading and surrounding the tumor. We observed that imiquimod treatment leads either to complete resolution or to a significant reduction of the tumors. Indeed, reduced proliferation and increased apoptosis were observed in tumors treated with imiquimod, and similar observations have recently been reported (56). Surprisingly, we found that the number of CD8+ T cells was much higher in vehicle-treated tumors, and few of these cells could be detected after imiquimod treatment regardless of the clinical response of the tumor to the drug. However, CD4+CD3− cells as well as CD4+MHC-II+, CD4+GR-1+, and GR-1+MHC-II+ were abundantly present in all imiquimod-treated tumors, and their numerical increase was more substantial in complete responders and in mice with stable disease. Because the number of these cells directly correlated with the clinical response to the drug, we propose that the antitumor property of imiquimod is due in part to the recruitment of pDC-like cells. Indeed, pDCs represent the highest type I IFN-producing cells in both humans and mice. IFN-α has been used extensively in the treatment of metastatic melanoma, mediating the regression of advanced disease in 15–20% of patients when used at high doses (57, 58). Despite these findings, it remains unclear whether IFN-α mediates its antitumor effects through stimulation of the host’s immune system, by a direct effect on the tumor cells, or both (59). IFN-α can exert direct effects on tumor cells because it is a potent inhibitor of cell proliferation in vitro, and it strongly up-regulates the expression of MHC-I Ags and adhesion molecules such as ICAM-1 and L-selectin on tumor cells (60, 61). It has also been demonstrated that IFN-α effectively inhibits the release of tumor-derived, proangiogenic factors such as basic fibroblast growth factor (62, 63). Interestingly, we found that the expression of IFN-α was increased in spleen cells after imiquimod treatment. According to our findings and on the basis of previous results, we propose that the antitumor properties of imiquimod are related to the recruitment of pDCs and to the release of IFN-α by these cells. In conclusion, this study provides a detailed and exhaustive view of the cellular composition of the inflammatory infiltrate induced by imiquimod in normal and neoplastic skin and suggest that the immunostimulatory and tumoricidal effects of the drug might be mediated at least in part by a specific subtype of DCs, termed pDC-like cells.

Acknowledgments

We are grateful to Martina Hammer for maintaining our mouse colonies, to Dr. Dieter Maurer for help with FACS analysis, and to Dr. Caterina Barresi for constant support. Imiquimod cream (5%; Aldara) was a generous gift from Werner N. Peljak (Pelpharma, Vienna, Austria) and Olivier Loudon (3M Pharmaceuticals, Austria). The Aldara vehicle was provided by Dr. Eggert Stockfleth (3M Pharmaceuticals, Perchtolsdorf, Austria). We thank Drs. Antonio Costanzo, José M. Carballido, Dieter Maurer, and Erwin F. Wagner for critical reading of the manuscript.

Footnotes

  • The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

  • ↵1 This work was supported by the Competence Center for Biomolecular Therapeutics, funded by the Austrian Ministry of Science and Technology, City of Vienna, and industrial partners.

  • ↵2 Current address: Department of Dermatology, University of Rome Tor Vergata, Viale Oxford 81, 00133 Rome, Italy.

  • ↵3 Address correspondence and reprint requests to Dr. Maria Sibilia, Department of Dermatology, Division of Immunology, Allergy and Infectious Diseases, Medical University of Vienna and Competence Center for Biomolecular Therapeutics, Vienna Competence Center, Lazarettgasse 19, A-1090 Vienna, Austria. E-mail address: maria.sibilia{at}meduniwien.ac.at

  • ↵4 Abbreviations used in this paper: DC, dendritic cell; pDC, plasmacytoid DC; LC, Langerhans cell; LN, lymph node; MHC-II, MHC class II.

  • Received March 22, 2004.
  • Accepted June 30, 2004.

References

  1. ↵
    Banchereau, J., F. Briere, C. Caux, J. Davoust, S. Lebecque, Y. J. Liu, B. Pulendran, K. Palucka. 2000. Immunobiology of dendritic cells. Annu. Rev. Immunol. 18:767.
    OpenUrlCrossRefPubMed
  2. ↵
    Mellman, I., R. M. Steinman. 2001. Dendritic cells: specialized and regulated antigen processing machines. Cell 106:255.
    OpenUrlCrossRefPubMed
  3. ↵
    Ardavin, C., G. Martinez del Hoyo, P. Martin, F. Anjuere, C. F. Arias, A. R. Marin, S. Ruiz, V. Parrillas, H. Hernandez. 2001. Origin and differentiation of dendritic cells. Trends Immunol. 22:691.
    OpenUrlCrossRefPubMed
  4. ↵
    Vandenabeele, S., L. Wu. 1999. Dendritic cell origins: puzzles and paradoxes. Immunol. Cell. Biol. 77:411.
    OpenUrlCrossRefPubMed
  5. ↵
    Shortman, K., Y. J. Liu. 2002. Mouse and human dendritic cell subtypes. Nat. Rev. Immunol. 2:151.
    OpenUrlCrossRefPubMed
  6. ↵
    Wollenberg, A., M. Wagner, S. Gunther, A. Towarowski, E. Tuma, M. Moderer, S. Rothenfusser, S. Wetzel, S. Endres, G. Hartmann. 2002. Plasmacytoid dendritic cells: a new cutaneous dendritic cell subset with distinct role in inflammatory skin diseases. J. Invest. Dermatol. 119:1096.
    OpenUrlCrossRefPubMed
  7. ↵
    Bangert, C., J. Friedl, G. Stary, G. Stingl, T. Kopp. 2003. Immunopathologic features of allergic contact dermatitis in humans: participation of plasmacytoid dendritic cells in the pathogenesis of the disease?. J. Invest. Dermatol. 121:1409.
    OpenUrlCrossRefPubMed
  8. ↵
    Lennert, K., W. Remmele. 1958. Karyometrische Untersuchungen an Lymphknotenzellen des Menschen. I. Mitt Germinoblasten, Lymphoblasten und Lymphozyten. Acta Haematol. 19:99.
    OpenUrlCrossRefPubMed
  9. ↵
    Kohrguber, N., N. Halanek, M. Gröger, D. Winter, K. Rappersberger, M. Schmitt-Egenolf, G. Stingl, D. Maurer. 1999. Survival, maturation, and function of CD11c− and CD11c+ peripheral blood dendritic cells are differentially regulated by cytokines. J. Immunol. 163:3250.
    OpenUrlAbstract/FREE Full Text
  10. ↵
    Cella, M., D. Jarrossay, F. Facchetti, O. Alebardi, H. Nakajima, A. Lanzavecchia, M. Colonna. 1999. Plasmacytoid monocytes migrate to inflamed lymph nodes and produce large amounts of type I interferon. Nat. Med. 5:919.
    OpenUrlCrossRefPubMed
  11. ↵
    Siegal, F. P., N. Kadowaki, M. Shodell, P. A. Fitzgerald-Bocarsly, K. Shah, S. Ho, S. Antonenko, Y. J. Liu. 1999. The nature of the principal type 1 interferon-producing cells in human blood. Science 284:1835.
    OpenUrlAbstract/FREE Full Text
  12. ↵
    Hochrein, H., M. O’Keeffe, H. Wagner. 2002. Human and mouse plasmacytoid dendritic cells. Hum. Immunol. 63:1103.
    OpenUrlCrossRefPubMed
  13. ↵
    Galy, A., I. Christopherson, G. Ferlazzo, G. Liu, H. Spits, K. Georgopoulos. 2000. Distinct signals control the hematopoiesis of lymphoid-related dendritic cells. Blood 95:128.
    OpenUrlAbstract/FREE Full Text
  14. ↵
    Abb, J., H. Abb, F. Deinhardt. 1983. Phenotype of human α-interferon producing leucocytes identified by monoclonal antibodies. Clin. Exp. Immunol. 52:179.
    OpenUrlPubMed
  15. ↵
    Kadowaki, N., S. Antonenko, J. Y. Lau, Y. J. Liu. 2000. Natural interferon α/β-producing cells link innate and adaptive immunity. J. Exp. Med. 192:219.
    OpenUrlAbstract/FREE Full Text
  16. ↵
    Asselin-Paturel, C., A. Boonstra, M. Dalod, I. Durand, N. Yessaad, C. Dezutter-Dambuyant, A. Vicari, A. O’Garra, C. Biron, F. Briere, et al 2001. Mouse type I IFN-producing cells are immature APCs with plasmacytoid morphology. Nat. Immunol. 2:1144.
    OpenUrlCrossRefPubMed
  17. ↵
    Bjorck, P.. 2001. Isolation and characterization of plasmacytoid dendritic cells from Flt3 ligand and granulocyte-macrophage colony-stimulating factor-treated mice. Blood 98:3520.
    OpenUrlAbstract/FREE Full Text
  18. ↵
    Nakano, H., M. Yanagita, M. D. Gunn. 2001. CD11c+B220+Gr-1+ cells in mouse lymph nodes and spleen display characteristics of plasmacytoid dendritic cells. J. Exp. Med. 194:1171.
    OpenUrlAbstract/FREE Full Text
  19. ↵
    Nikolic, T., G. M. Dingjan, P. J. Leenen, R. W. Hendriks. 2002. A subfraction of B220+ cells in murine bone marrow and spleen does not belong to the B cell lineage but has dendritic cell characteristics. Eur. J. Immunol. 32:686.
    OpenUrlCrossRefPubMed
  20. ↵
    Ferrero, I., W. Held, A. Wilson, F. Tacchini-Cottier, F. Radtke, H. R. MacDonald. 2002. Mouse CD11c+ B220+ Gr1+ plasmacytoid dendritic cells develop independently of the T-cell lineage. Blood 100:2852.
    OpenUrlAbstract/FREE Full Text
  21. ↵
    Martin, P., G. M. Del Hoyo, F. Anjuere, C. F. Arias, H. H. Vargas, L. A. Fernandez, V. Parrillas, C. Ardavin. 2002. Characterization of a new subpopulation of mouse CD8α+ B220+ dendritic cells endowed with type 1 interferon production capacity and tolerogenic potential. Blood 100:383.
    OpenUrlAbstract/FREE Full Text
  22. ↵
    O’Keeffe, M., H. Hochrein, D. Vremec, I. Caminschi, J. L. Miller, E. M. Anders, L. Wu, M. H. Lahoud, S. Henri, B. Scott, et al 2002. Mouse plasmacytoid cells: long-lived cells, heterogeneous in surface phenotype and function, that differentiate into CD8+ dendritic cells only after microbial stimulus. J. Exp. Med. 196:1307.
    OpenUrlAbstract/FREE Full Text
  23. ↵
    Vremec, D., J. Pooley, H. Hochrein, L. Wu, K. Shortman. 2000. CD4 and CD8 expression by dendritic cell subtypes in mouse thymus and spleen. J. Immunol. 164:2978.
    OpenUrlAbstract/FREE Full Text
  24. ↵
    Okada, T., Z. X. Lian, M. Naiki, A. A. Ansari, S. Ikehara, M. E. Gershwin. 2003. Murine thymic plasmacytoid dendritic cells. Eur. J. Immunol. 33:1012.
    OpenUrlCrossRefPubMed
  25. ↵
    Lian, Z. X., T. Okada, X. S. He, H. Kita, Y. J. Liu, A. A. Ansari, K. Kikuchi, S. Ikehara, M. E. Gershwin. 2003. Heterogeneity of dendritic cells in the mouse liver: identification and characterization of four distinct populations. J. Immunol. 170:2323.
    OpenUrlAbstract/FREE Full Text
  26. ↵
    Akira, S., K. Takeda, T. Kaisho. 2001. Toll-like receptors: critical proteins linking innate and acquired immunity. Nat. Immunol. 2:675.
    OpenUrlCrossRefPubMed
  27. ↵
    Akira, S., H. Hemmi. 2003. Recognition of pathogen-associated molecular patterns by TLR family. Immunol. Lett. 85:85.
    OpenUrlCrossRefPubMed
  28. ↵
    Gibson, S. J., J. M. Lindh, T. R. Riter, R. M. Gleason, L. M. Rogers, A. E. Fuller, J. L. Oesterich, K. B. Gorden, X. Qiu, S. W. McKane, et al 2002. Plasmacytoid dendritic cells produce cytokines and mature in response to the TLR7 agonists, imiquimod and resiquimod. Cell. Immunol. 218:74.
    OpenUrlCrossRefPubMed
  29. ↵
    Hemmi, H., T. Kaisho, O. Takeuchi, S. Sato, H. Sanjo, K. Hoshino, T. Horiuchi, H. Tomizawa, K. Takeda, S. Akira. 2002. Small anti-viral compounds activate immune cells via the TLR7 MyD88-dependent signaling pathway. Nat. Immunol. 3:196.
    OpenUrlCrossRefPubMed
  30. ↵
    Heil, F., H. Hemmi, H. Hochrein, F. Ampenberger, C. Kirschning, S. Akira, G. Lipford, H. Wagner, S. Bauer. 2004. Species-specific recognition of single-stranded RNA via Toll-like receptor 7 and 8. Science 303:1526.
    OpenUrlAbstract/FREE Full Text
  31. ↵
    Diebold, S. S., T. Kaisho, H. Hemmi, S. Akira, E. S. C. Reis. 2004. Innate antiviral responses by means of TLR7-mediated recognition of single-stranded RNA. Science 303:1529.
    OpenUrlAbstract/FREE Full Text
  32. ↵
    Stanley, M. A.. 2002. Imiquimod and the imidazoquinolones: mechanism of action and therapeutic potential. Clin. Exp. Dermatol. 27:571.
    OpenUrlCrossRefPubMed
  33. ↵
    Miller, R. L., J. F. Gerster, M. L. Owens, H. B. Slade, M. A. Tomai. 1999. Imiquimod applied topically: a novel immune response modifier and new class of drug. Int. J. Immunopharmacol. 21:1.
    OpenUrlCrossRefPubMed
  34. ↵
    Rhodes, J.. 2002. Discovery of immunopotentiatory drugs: current and future strategies. Clin. Exp. Immunol. 130:363.
    OpenUrlCrossRefPubMed
  35. ↵
    Megyeri, K., W. C. Au, I. Rosztoczy, N. B. Raj, R. L. Miller, M. A. Tomai, P. M. Pitha. 1995. Stimulation of interferon and cytokine gene expression by imiquimod and stimulation by Sendai virus utilize similar signal transduction pathways. Mol. Cell. Biol. 15:2207.
    OpenUrlAbstract/FREE Full Text
  36. ↵
    Kono, T., S. Kondo, S. Pastore, G. M. Shivji, M. A. Tomai, R. C. McKenzie, D. N. Sauder. 1994. Effects of a novel topical immunomodulator, imiquimod, on keratinocyte cytokine gene expression. Lymphokine Cytokine Res. 13:71.
    OpenUrlPubMed
  37. ↵
    Ito, T., R. Amakawa, T. Kaisho, H. Hemmi, K. Tajima, K. Uehira, Y. Ozaki, H. Tomizawa, S. Akira, S. Fukuhara. 2002. Interferon-α and interleukin-12 are induced differentially by Toll-like receptor 7 ligands in human blood dendritic cell subsets. J. Exp. Med. 195:1507.
    OpenUrlAbstract/FREE Full Text
  38. ↵
    Testerman, T. L., J. F. Gerster, L. M. Imbertson, M. J. Reiter, R. L. Miller, S. J. Gibson, T. L. Wagner, M. A. Tomai. 1995. Cytokine induction by the immunomodulators imiquimod and S-27609. J. Leukocyte Biol. 58:365.
    OpenUrlAbstract
  39. ↵
    Harrison, C. J., R. L. Miller, D. I. Bernstein. 1994. Posttherapy suppression of genital herpes simplex virus (HSV) recurrences and enhancement of HSV-specific T-cell memory by imiquimod in guinea pigs. Antimicro.b Agents Chemother. 38:2059.
    OpenUrlAbstract/FREE Full Text
  40. ↵
    Harrison, C. J., R. L. Miller, D. I. Bernstein. 2001. Reduction of recurrent HSV disease using imiquimod alone or combined with a glycoprotein vaccine. Vaccine 19:1820.
    OpenUrlCrossRefPubMed
  41. ↵
    Sidky, Y., G. Bryan, J. Weeks, J. Hatcher, E. Borden. 1990. Effect of treatment with an oral interferon inducer, imidazoquinolamine (R-837) on the growth of mouse bladder carcinoma FCB. J Interferon Res. 10:S123. (Abstr.).
    OpenUrl
  42. ↵
    Sidky, Y. A., E. C. Borden, C. E. Weeks, M. J. Reiter, J. F. Hatcher, G. T. Bryan. 1992. Inhibition of murine tumor growth by an interferon-inducing imidazoquinolinamine. Cancer Res. 52:3528.
    OpenUrlAbstract/FREE Full Text
  43. ↵
    Suzuki, H., B. Wang, G. M. Shivji, P. Toto, P. Amerio, M. A. Tomai, R. L. Miller, D. N. Sauder. 2000. Imiquimod, a topical immune response modifier, induces migration of Langerhans cells. J. Invest. Dermatol. 114:135.
    OpenUrlCrossRefPubMed
  44. ↵
    Tomai, M. A., S. J. Gibson, L. M. Imbertson, R. L. Miller, P. E. Myhre, M. J. Reiter, T. L. Wagner, C. B. Tamulinas, J. M. Beaurline, J. F. Gerster, et al 1995. Immunomodulating and antiviral activities of the imidazoquinoline S-28463. Antiviral Res. 28:253.
    OpenUrlCrossRefPubMed
  45. ↵
    Wagner, S. N., C. Wagner, P. Luehrs, T. K. Weimann, R. Kutil, M. Goos, G. Stingl, A. Schneeberger. 2000. Intracutaneous genetic immunization with autologous melanoma-associated antigen Pmel17/gp100 induces T cell-mediated tumor protection in vivo. J. Invest. Dermatol. 115:1082.
    OpenUrlCrossRefPubMed
  46. ↵
    Wang, B., A. Rieger, O. Kilgus, K. Ochiai, D. Maurer, D. Foedinger, J. P. Kinet, G. Stingl. 1992. Epidermal Langerhans cells from normal human skin bind monomeric IgE via FcεRI. J. Exp. Med. 175:1353.
    OpenUrlAbstract/FREE Full Text
  47. ↵
    Montoya, M., G. Schiavoni, F. Mattei, I. Gresser, F. Belardelli, P. Borrow, D. F. Tough. 2002. Type I interferons produced by dendritic cells promote their phenotypic and functional activation. Blood 99:3263.
    OpenUrlAbstract/FREE Full Text
  48. ↵
    Bottcher, T., M. von Mering, S. Ebert, U. Meyding-Lamade, U. Kuhnt, J. Gerber, R. Nau. 2003. Differential regulation of Toll-like receptor mRNAs in experimental murine central nervous system infections. Neurosci. Lett. 344:17.
    OpenUrlCrossRefPubMed
  49. ↵
    Spergel, J. M., E. Mizoguchi, J. P. Brewer, T. R. Martin, A. K. Bhan, R. S. Geha. 1998. Epicutaneous sensitization with protein antigen induces localized allergic dermatitis and hyperresponsiveness to methacholine after single exposure to aerosolized antigen in mice. J. Clin. Invest. 101:1614.
    OpenUrlCrossRefPubMed
  50. ↵
    Edwards, A. D., S. S. Diebold, E. M. Slack, H. Tomizawa, H. Hemmi, T. Kaisho, S. Akira, C. Reis e Sousa. 2003. Toll-like receptor expression in murine DC subsets: lack of TLR7 expression by CD8α+ DC correlates with unresponsiveness to imidazoquinolines. Eur. J. Immunol. 33:827.
    OpenUrlCrossRefPubMed
  51. ↵
    Steinmann, A., J. O. Funk, G. Schuler, P. von den Driesch. 2000. Topical imiquimod treatment of a cutaneous melanoma metastasis. J. Am. Acad. Dermatol. 43:555.
    OpenUrlCrossRefPubMed
  52. ↵
    Ahmed, I., J. Berth-Jones. 2000. Imiquimod: a novel treatment for lentigo maligna. Br. J. Dermatol. 143:843.
    OpenUrlCrossRefPubMed
  53. ↵
    Hengge, U. R., R. Stark. 2001. Topical imiquimod to treat intraepidermal carcinoma. Arch. Dermatol. 137:709.
    OpenUrlPubMed
  54. ↵
    Imbertson, L. M., J. M. Beaurline, A. M. Couture, S. J. Gibson, R. M. Smith, R. L. Miller, M. J. Reiter, T. L. Wagner, M. A. Tomai. 1998. Cytokine induction in hairless mouse and rat skin after topical application of the immune response modifiers imiquimod and S-28463. J. Invest. Dermatol. 110:734.
    OpenUrlCrossRefPubMed
  55. ↵
    Roake, J. A., A. S. Rao, P. J. Morris, C. P. Larsen, D. F. Hankins, J. M. Austyn. 1995. Dendritic cell loss from nonlymphoid tissues after systemic administration of lipopolysaccharide, tumor necrosis factor, and interleukin 1. J. Exp. Med. 181:2237.
    OpenUrlAbstract/FREE Full Text
  56. ↵
    Sidbury, R., N. Neuschler, E. Neuschler, P. Sun, X. Q. Wang, R. Miller, M. Tomai, E. Puscasiu, S. Gugneja, A. S. Paller. 2003. Topically applied imiquimod inhibits vascular tumor growth in vivo. J. Invest. Dermatol. 121:1205.
    OpenUrlCrossRefPubMed
  57. ↵
    Kirkwood, J. M., J. G. Ibrahim, V. K. Sondak, M. S. Ernstoff, M. Ross. 2002. Interferon α-2a for melanoma metastases. Lancet 359:978.
    OpenUrlCrossRefPubMed
  58. ↵
    Belardelli, F., M. Ferrantini, E. Proietti, J. M. Kirkwood. 2002. Interferon-α in tumor immunity and immunotherapy. Cytokine Growth Factor Rev. 13:119.
    OpenUrlCrossRefPubMed
  59. ↵
    Agarwala, S. S., J. M. Kirkwood. 1995. Potential uses of interferon α2 as adjuvant therapy in cancer. Ann. Surg. Oncol. 2:365.
    OpenUrlCrossRefPubMed
  60. ↵
    Martin-Henao, G. A., R. Quiroga, A. Sureda, J. R. Gonzalez, V. Moreno, J. Garcia. 2000. L-selectin expression is low on CD34+ cells from patients with chronic myeloid leukemia and interferon-α up-regulates this expression. Haematologica 85:139.
    OpenUrlPubMed
  61. ↵
    von Stamm, U., E. B. Brocker, M. von Depka Prondzinski, D. J. Ruiter, P. Rumke, C. Broding, S. Carrel, F. J. Lejeune. 1993. Effects of systemic interferon-α (IFN-α) on the antigenic phenotype of melanoma metastases: EORTC melanoma group cooperative study no. 18852. Melonoma Res. 3:173.
    OpenUrlCrossRef
  62. ↵
    Slaton, J. W., P. Perrotte, K. Inoue, C. P. Dinney, I. J. Fidler. 1999. Interferon-α-mediated down-regulation of angiogenesis-related genes and therapy of bladder cancer are dependent on optimization of biological dose and schedule. Clin. Cancer Res. 5:2726.
    OpenUrlAbstract/FREE Full Text
  63. ↵
    Ezekowitz, R. A., J. B. Mulliken, J. Folkman. 1992. Interferon α-2a therapy for life-threatening hemangiomas of infancy. N. Engl. J. Med. 326:1456.
    OpenUrlCrossRefPubMed
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