Cycle length: 28 days Dosage: 200-600 μg/kg 200-600 μg/2.2 lb Day 1-7 200 μg/2.2 lb Day 8-14 400 μg/2.2 lb Day 15-21 600 μg/2.2 lb Day 22-28 200 μg/2.2 lb Based on the available evidence, there is no established or FDA-approved dose of ivermectin for cancer treatment, as ivermectin is currently being investigated as a repositioned drug for cancer with doses of 200-600 μg/kg in preclinical studies. Preclinical Evidence for Ivermectin in Cancer Dosing in Laboratory Studies In preclinical cancer research, ivermectin has been studied at concentrations of approximately 5 μM in cell culture studies 1 This concentration showed antitumor effects in several cancer cell lines, with breast cancer (MDA-MB-231, MDA-MB-468, MCF-7) and ovarian cancer (SKOV-3) being most sensitive Prostate cancer cell line DU145 was found to be most resistant to ivermectin treatment 1 Mechanisms of Action in Cancer Ivermectin has demonstrated potential anticancer effects through: Cell cycle arrest at G0-G1 phase with modulation of cell cycle control proteins 1 Inhibition of tumor metastasis via suppression of the Wnt/β-catenin/integrin β1/FAK signaling pathway 2 Synergistic effects when combined with conventional chemotherapeutics like docetaxel, cyclophosphamide, and tamoxifen 1 Human Dosing Considerations Standard Ivermectin Dosing for Approved Indications For FDA-approved indications (onchocerciasis, strongyloidiasis), ivermectin is typically administered at 200 μg/kg orally 3 Pharmacokinetic studies show that: Peak plasma concentrations occur approximately 4 hours after dosing Plasma half-life is approximately 18 hours following oral administration Bioavailability increases 2.5-fold when taken with a high-fat meal 3 Safety of Higher Doses Animal studies have investigated higher doses of ivermectin (0.22-170 mg/kg) using special polyelectrolyte formulations 4 These higher doses showed no apparent damage to intestinal tissues in the animal model, though some delocalization of Myosin-Vb at enterocytes was observed at the highest dose Ivermectin (Systemic Adult Medication This information from Lexicomp® explains what you need to know about this medication, including what it’s used for, how to take it, its side effects, and when to call your healthcare provider. Brand Names: US Stromectol Brand Names: Canada Stromectol What is this drug used for? It is used to treat certain parasite infections. What do I need to tell my doctor BEFORE I take this drug? If you are allergic to this drug; any part of this drug; or any other drugs, foods, or substances. Tell your doctor about the allergy and what signs you had. This drug may interact with other drugs or health problems. Tell your doctor and pharmacist about all of your drugs (prescription or OTC, natural products, vitamins) and health problems. You must check to make sure that it is safe for you to take this drug with all of your drugs and health problems. Do not start, stop, or change the dose of any drug without checking with your doctor. What are some things I need to know or do while I take this drug? Tell all of your health care providers that you take this drug. This includes your doctors, nurses, pharmacists, and dentists. You will need to have a stool exam after taking this drug to make sure the infection is gone. Some patients may need to take another dose of this drug to treat the infection. Talk with the doctor. To lower the chance of feeling dizzy or passing out, rise slowly if you have been sitting or lying down. Be careful going up and down stairs. Tell your doctor if you are pregnant, plan on getting pregnant, or are breast-feeding. You will need to talk about the benefits and risks to you and the baby. What are some side effects that I need to call my doctor about right away? WARNING/CAUTION: Even though it may be rare, some people may have very bad and sometimes deadly side effects when taking a drug. Tell your doctor or get medical help right away if you have any of the following signs or symptoms that may be related to a very bad side effect: Signs of an allergic reaction, like rash; hives; itching; red, swollen, blistered, or peeling skin with or without fever; wheezing; tightness in the chest or throat; trouble breathing, swallowing, or talking; unusual hoarseness; or swelling of the mouth, face, lips, tongue, or throat. Severe dizziness or passing out. Change in eyesight, eye pain, or severe eye irritation. Joint pain. Swelling. Fever. Swollen gland. Nervous system problems have happened with this drug. Sometimes, this may even lead to coma or death. Call your doctor right away if you feel very sleepy, sluggish, confused, or out of sorts. A very bad and sometimes deadly brain problem has happened in some patients who also have Loa loa infection. This is rare. Tell your doctor if you have been to West Africa or Central Africa where you can get this infection. Tell your doctor right away if you have signs like neck pain, back pain, red eyes, eye bleeding, shortness of breath, seizures, trouble walking or standing, confusion, feeling very tired or weak, or if you are unable to control your bladder or bowels. What are some other side effects of this drug? All drugs may cause side effects. However, many people have no side effects or only have minor side effects. Call your doctor or get medical help if any of these side effects or any other side effects bother you or do not go away: Itching. Skin irritation. These are not all of the side effects that may occur. If you have questions about side effects, call your doctor. Call your doctor for medical advice about side effects. You may report side effects to your national health agency. You may report side effects to the FDA at 1-800-332-1088. You may also report side effects at https://www.fda.gov/medwatch. How is this drug best taken? Use this drug as ordered by your doctor. Read all information given to you. Follow all instructions closely. Take on an empty stomach. Take with a full glass of water. Drink lots of noncaffeine liquids unless told to drink less liquid by your doctor. What do I do if I miss a dose? Only 1 dose of this drug is needed. If you miss your dose, take it as soon as you think about it. How do I store and/or throw out this drug? Store at room temperature in a dry place. Do not store in a bathroom. Keep all drugs in a safe place. Keep all drugs out of the reach of children and pets. Throw away unused or expired drugs. Do not flush down a toilet or pour down a drain unless you are told to do so. Check with your pharmacist if you have questions about the best way to throw out drugs. There may be drug take-back programs in your area. General drug facts If your symptoms or health problems do not get better or if they become worse, call your doctor. Do not share your drugs with others and do not take anyone else’s drugs. Some drugs may have another patient information leaflet. If you have any questions about this drug, please talk with your doctor, nurse, pharmacist, or other health care provider. Some drugs may have another patient information leaflet. Check with your pharmacist. If you have any questions about this drug, please talk with your doctor, nurse, pharmacist, or other health care provider. If you think there has been an overdose, call your poison control center or get medical care right away. Be ready to tell or show what was taken, how much, and when it happened. Consumer Information Use and Disclaimer This generalized information is a limited summary of diagnosis, treatment, and/or medication information. It is not meant to be comprehensive and should be used as a tool to help the user understand and/or assess potential diagnostic and treatment options. 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Checkpoint blockade1,2 has emerged as a revolutionary approach that harnesses a patient’s own immune system to treat cancer. However, checkpoint inhibitors as single agents are only effective in a subset of patients and cancer types2. Recent studies suggest that efficacy of checkpoint inhibitors is primarily limited to cancers already infiltrated by T cells—often termed “hot” tumors. In contrast, “cold” tumors have little to no T cell infiltration and generally do not respond to checkpoint blockade. Clinical studies with checkpoint blockade therapy in breast cancer have focused on triple negative breast cancer (TNBC), because this subtype has a higher mutational load and is thought to be more “immunogenic”3. While an early phase Ib study (KEYNOTE-012) of pembrolizumab (anti-PD1 antibody) monotherapy showed preliminary evidence of clinical activity in a small subset (18.5%) of advanced TNBC patients4, the phase 3 study (KEYNOTE-119) showed no improvement over chemotherapy5. Beyond monotherapy, checkpoint blockade plus chemotherapy combinations are being investigated. Atezolizumab (anti-PDL1 antibody) in combination with nab-paclitaxel demonstrated efficacy for PD-L1-positive unresectable locally advanced or metastatic TNBC in IMpassion130 (ref. 6), leading to the first FDA approval of immune checkpoint therapy for breast cancer in March 2019. However, IMpassion131 (atezolizumab + paclitaxel) was recently reported to be negative7. These results highlight the continued challenge of breast cancer for immune checkpoint therapies. As such, there is considerable need to identify drugs capable of priming breast tumors (turning “cold” tumors “hot”) to synergize with checkpoint blockade. A recently described phenomenon, termed immunogenic cell death (ICD)8,9, is a form of cell death that induces an immune response from the host. ICD is distinguished from classical apoptosis and other non-immunogenic or tolerogenic forms of cell death by several hallmarks, including release of adenosine triphosphate (ATP) and high-mobility group box 1 protein (HMGB1), and surface exposure of calreticulin8,9,10. In cancer patients, ICD-based anti-tumor immune responses are linked to beneficial outcomes produced by some conventional chemotherapeutic agents11,12,13,14. For example, efficacy of anthracyclines in breast cancer15,16,17 and oxaliplatin in colorectal cancer18 correlates with post-treatment increases in the ratio of cytotoxic CD8+ T lymphocytes to FoxP3+ regulatory T cells within the tumor. In contrast, poor responses to chemotherapy in solid tumors are associated with lymphopenia19. Thus, ICD-inducing chemotherapy appears to work in conjunction with the host immune system to achieve efficacy. However, chemotherapy is a double-edged sword: it can suppress as well as stimulate immune cells. An agent that induces ICD of cancer cells without suppressing immune function would be ideal for combination with checkpoint blockade. Seeking such an agent among FDA-approved drugs, our group found that the anti-parasitic agent ivermectin promotes ICD in breast cancer cells20. Among our previous findings was evidence that ivermectin, an anti-parasitic drug used worldwide since 1975, modulates the P2X4/P2X7 purinergic pathway, suggesting that ivermectin may further harness tumors’ intrinsic high extracellular levels of ATP for anti-cancer activity. Of note, P2X purinoceptor 4 and P2X purinoceptor 7 (P2X4/P2X7) are widely expressed on various immune subpopulations, suggesting that ivermectin might also have direct immunomodulatory effects. Results Ivermectin can turn “cold” breast tumors “hot” We studied the effects of ivermectin in vivo using the 4T1 mouse model of TNBC. HMGB1 is a chromatin protein present in all cells and its release is a hallmark of ICD21. HMGB1 staining (green) was observed uniformly across the entire tumor from untreated mice (Fig. 1A). In contrast, tumors isolated from mice treated with ivermectin showed large areas of DAPI-positive cells lacking HMGB1 (Fig. 1B), suggesting that HMGB1 had been released into the extracellular space. Ivermectin treatment also altered calreticulin expression, with higher levels (green) observed in tumors from treated animals, indicating a significant increase in this ICD-associated prophagocytic signal and mediator (Fig. 1C, D). Exposure of endosomal calreticulin onto the surface of ER-stressed, damaged or dying cells promotes immunogenic phagocytosis and antigen cross-presentation by antagonizing both the “don’t eat me” signals associated with CD47 and the tolerogenic “eat me” signals associated with phosphatidylserine (PS) exposure, while promoting the interaction with its receptor low-density lipoprotein-receptor related protein (LRP) on phagocytic cells. Robust infiltration of both CD4+ and CD8+ T cells was seen in ivermectin-treated tumors (Fig. 1F) but not in untreated tumors (Fig. 1E). Significantly higher percentages of cells were positive for CD4 (p < 0.01, Fig. 1G) and CD8 (p < 0.0001, Fig. 1H) in ivermectin-treated than in untreated tumors. Together, these data indicate that treatment with ivermectin induced hallmarks of ICD within 4T1 breast tumors and recruited large numbers of CD4+ and CD8+ T cells into these tumors. To further confirm that ivermectin induces ICD in vivo, we also utilized a classical vaccination approach considered as the gold standard for detection of ICD: treatment of 4T1 cells with IVM to induce ICD in vitro followed by inoculation into naïve mice, then subsequent challenge with live 4T1 cells to demonstrate prevention of tumor outgrowth21. This experiment informed a possible induction of bona fide ICD by demonstrating protection against subsequent challenge with live 4T1 cells (p < 0.01, Fig. 1I). Fig. 1: Treatment with ivermectin induces immunogenic cell death (ICD) in vivo and recruitment of T cells into tumors. figure 1 4T1 breast tumors were isolated from mice that were untreated (left panels) or ivermectin-treated (right panels) daily for 14 days. Panels A, B show staining for HMGB1 (green), a hallmark of ICD. Panels C, D show staining for calreticulin (green), another hallmark of ICD. Staining for CK7 (red) identifies 4T1 cells. Data are representative of two independent experiments. Panels E, F show staining for CD4+ (green), CD8+ T cells (yellow), and cancer cells via staining for CK7 (red). Data are representative of three independent experiments. Panels G, H display quantitative data on T cell infiltration shown in E, F. Data were obtained by quantifying five random fields from whole tumor images. Panel I demonstrates the protective effect of prophylactic subcutaneous vaccination with 1 million 4T1 cells treated with 12 μM ivermectin ex vivo (24 h), then challenged contralaterally with live 4T1 cells 1 week post vaccination (n = 4). Statistical significance was evaluated using the linear mixed effects model of log tumor volume; *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001, ****p ≤ 0.0001. Full size image Direct immunomodulatory effects of ivermectin Ivermectin treatment in vivo did not produce any significant changes in the frequencies of various effector and regulatory CD4 (Fig. S2A) or CD8 (Fig. S2B) T cell subpopulations isolated from the spleens of treated animals. However, functional interrogation of splenocytes isolated from control vs. 4T1 tumor-bearing mice revealed significant immunomodulatory effects. Tumor-bearing mice 1 month post-inoculation developed enlarged spleens with an expanded population of CD11b+ myeloid cells (Fig. 2A), which includes both CD11b+GR-1+ myeloid-derived suppressor cells (MDSCs) and CD11b+GR-1− Monocytes/Macrophages (Mon/Mac). Ivermectin treatment ex vivo preferentially depleted this expanded CD11b+ myeloid population, normalizing the balance between myeloid and T cell compartments (Fig. 2A). Myeloid and lymphoid cell populations showed differential sensitivity to increasing doses of ivermectin (Fig. 2B and S2C). A linear mixed effects model of log cell count adjusted for cell type revealed that CD11b+ myeloid cells were the most sensitive to ivermectin, showing significant reductions with as little as 4 μM after 48 h, 8 μM after 24 h, or 16 μM after 4 h—demonstrating rapid and selective targeting of this immunosuppressive population (each result, p < 0.0001). In contrast, achieving similar reductions in CD4 or CD8 T cells required higher doses and/or longer exposure to ivermectin: observed in CD8 T cells only after 48 h of 8 μM or 24 h of 16 μM, and in CD4 T cells only after the maximum exposure (48 h of 16 μM). Consistent with ivermectin being an allosteric modulator of the ATP/P2X4/P2X7 signaling axis which operates in both cancer and immune cells, differential sensitivity in myeloid cells was P2X7-dependent (Fig. 2C). P2X7 blockade with 10 μM KN62 reversed the ex vivo depletion of CD11b+GR-1+ MDSCs, CD11b+GR-1− Mon/Mac, and other immune subsets by ivermectin (p < 0.001). To mimic more physiologically relevant conditions of exposure, we also treated splenocytes with lower non-cytotoxic doses of ivermectin and observed that over extended exposure, ivermectin had a significant potentiating effect on phytohemagglutinin (PHA)-stimulated T cells and augmented the ratios of both CD8+ and CD4+ Teff/Tregs (Fig. 2D). PHA induces nonspecific T cell stimulation by binding to glycosylated surface proteins, including but not exclusive to the T cell receptor (TCR), causing TCR crosslinking and downstream signaling resulting in T cell activation and proliferation. The immuno-potentiating effects of extended exposure to lower non-cytotoxic doses of ivermectin were enhanced upon TCR stimulation (via PHA) and were inhibited in splenocytes from tumor-bearing mice (Fig. 2D), where different mechanisms including MDSCs as well as PD-1-mediated immunosuppression are known to interfere with proper TCR signaling and function. Fig. 2: Immunomodulatory effects of ivermectin ex vivo. figure 2 Splenocytes (SPL) were isolated from the spleens of aged-matched untreated and naïve non-tumor-bearing control mice (CTRL) or untreated 4T1 tumor bearing mice (TB), 1 month post tumor inoculation, then cultured on 96-well tissue culture-treated plates in complete R10 medium for 4h–48h and analyzed by flow cytometry for spontaneous and ivermectin-induced changes in various immune subpopulations. A Depletion of the expanded CD11b+ myeloid cells isolated from the spleens of tumor-bearing mice by ivermectin treatment ex vivo. B, C Splenocytes isolated from 4T1 tumor-bearing mice were exposed to increasing doses of ivermectin for 4 h or 48 h showing differential dose- and time-dependent sensitivity of different immune subpopulations (see also Fig. S2C). Depletion of CD11b+GR-1+ MDSCs, CD11b+GR-1− Monocytes/Macrophages, CD19+ B cells and CD3+ T cells by IVM could be reversed by an inhibitor of P2X7/CaMKII (KN62 at 10 μM). D Splenocytes from aged-matched untreated and naïve non-tumor-bearing control (CTRL) and 4T1 tumor-bearing (TB) mice were incubated for 24 h and 4 days with increasing doses of ivermectin (1–16 μM) with or without PHA to mimic TCR stimulation. Plots show averages and standard deviation based on triplicates; data representative of at least two independent experiments. Statistical significance versus (−) CTRL or as indicated was evaluated using the linear mixed effects model of log cell count adjusted for cell type: *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001, ****p ≤ 0.0001. Full size image Ivermectin synergizes with anti-PD1 antibody to control tumor growth and induces protective immunity The anti-cancer ICD and direct immunomodulatory effects of ivermectin raised the possibility that it could be combined with checkpoint blockade. We next investigated the efficacy of ivermectin and anti-PD1 antibody, alone or in combination, relative to no treatment (schema in Fig. S1A). Mean tumor volume over time was significantly decreased by the ivermectin and anti-PD1 antibody combination relative to no treatment (p < 0.001, Fig. 3A). Through a joint statistical model of longitudinal tumor volumes, ivermectin and anti-PD1 antibody demonstrated synergistic activity, defined as an effect that is significantly greater than the sum of the drugs’ individual effects (submodel p = 0.008, false discovery rate/FDR 3%, Table 1). Complete tumor regression was observed in 6/15 mice on the combination treatment, 1/20 on ivermectin alone, 1/10 on anti-PD1 antibody alone, and 0/25 on no treatment. Mice that resolved tumors on the ivermectin and anti-PD1 combination therapy were re-challenged with 100,000 4T1 cells in the contralateral mammary fat pads. All of these mice resisted development of new tumors (Fig. 3B), while control naïve animals all developed tumors (data not shown). This suggests that combined treatment with ivermectin and anti-PD1 induces protective anti-tumor immunity in complete responders. Fig. 3: Ivermectin synergizes with anti-PD1 therapy to control tumor growth in vivo. figure 3 Mice were inoculated with 100,000 4T1 cells 4 days before initiating therapy with ivermectin alone (n = 20), anti-PD1 antibody alone (n = 10), both drugs (n = 15), or no treatment (n = 25). A Tumor volume in control and treated animals; *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001. B Tumor growth in individual animals treated with ivermectin plus anti-PD1 antibody (five individual mice from one representative of three experiments shown). Three of five combination-treated animals completely resolved their tumors. Animals that resolved tumors were re-challenged with 100,000 4T1 cells on the contralateral mammary fat pad 30 days after the termination of therapy. Mice were observed and palpated twice a week for an additional 30 days for the establishment of a tumor mass. C–F Combination therapy with ivermectin and anti-PD1 recruits significantly more T cells into tumor sites and generates tumor-reactive CD8+ T cells. Tumors were isolated from mice at day 21. Staining was performed for nuclei (blue), CD4+ (green) cells, CD8+ cells (yellow), and tumor cells (red) (C). Percent positive for CD4 or CD8 was measured in five random fields in each group and divided by the number of nuclei in the field (D). Data are representative of two independent experiments; *p ≤ 0.05, **p ≤ 0.01, ****p ≤ 0.0001. Splenocytes isolated from tumor-bearing mice that received no treatment (n = 5), anti-PD1 alone (n = 5), or ivermectin with anti-PD1 (n = 4) were co-cultured with 4T1 cells. Reactive CD8+ cells were determined by CD107 mobilization and expression of IFNγ by flow cytometry. Representative flow plots for each treatment group are shown in E. F Percentage of CD8+ T cells reactive against 4T1 per mouse, grouped by treatment; **p ≤ 0.01. Full size image Table 1 Tumor growth in primary treatment. Full size table To gain further insight into the mechanism underlying efficacy of the combined treatment, we compared the magnitude to which ivermectin, anti-PD1, and their combination potentiated the infiltration of T cells. As shown visually in Fig. 3C and quantitatively in Fig. 3D, infiltration of both CD4+ and CD8+ T cells into 4T1 tumors (day 21) was greatest after treatment with the combination of ivermectin and anti-PD1. To measure anti-tumor T cells, splenocytes were isolated from untreated, single-agent-treated, or ivermectin plus anti-PD1 combination-treated mice, then co-cultured with 4T1 cells as targets to measure CD107 mobilization and IFN-γ expression as markers for functional T cell responses22. A functional tumor-specific immune response was confirmed by the presence of a discrete population of CD8+ T cells positive for CD107 and IFN-γ in mice treated with ivermectin plus anti-PD1, but not in mice treated with anti-PD1 alone or untreated controls (p < 0.01; Fig. 3E, F). Combination therapy effective across spectrum of clinically relevant settings Moving beyond control of primary tumors, we sought to test this combination immunotherapy across the major clinically relevant settings: neoadjuvant, adjuvant, and metastatic treatments. We also explored the effects of further augmenting this combination immunotherapy with interleukin-2 (IL-2). IL-2 was the first cytokine to be successfully used in the treatment of cancer to induce T cell activation23. A major challenge in the development of IL-2 as a therapeutic antitumor agent is that IL-2 can act on both T cells and regulatory T cells (Tregs). The contrasting actions of IL-2 has led to inconsistent responses and limited the development of high-dose IL-2 for cancer immunotherapy. Increasing the half-life of IL-2 has been shown to be a promising strategy for improving IL-2-based immunotherapy. This can greatly reduce the dose of IL-2 required for therapeutic activity, enhancing both safety and efficacy24,25. We explored the secondary hypothesis that addition of a recombinant albumin-IL-2 fusion with extended half-life to the ivermectin and anti-PD1 regimen (anti-PD-1 + IL-2 therapy, termed “IP” for simplicity) can further improve the efficacy of our combined treatment. Neoadjuvant therapy has come to play an increasingly prominent role in the treatment of cancer. We tested treatment of ivermectin combined with anti-PD-1 and IL-2 by monitoring survival of animals receiving neoadjuvant combination therapy followed by surgical resection of the primary tumor on day 16 following tumor inoculation (schema in Fig. S1A). Development of loco-regional recurrence and distant metastases were monitored by bioluminescent imaging, and animals were euthanized upon decline in body condition score and signs of morbidity. All untreated animals required euthanasia due to lethal diseases around day 20–25 following surgical resection of primary tumor (Fig. 4A). Treatment with IP therapy alone provided some survival benefit with ~40% of animals remaining free of lethal disease. The best survival outcome was seen with the combination of IP and ivermectin therapy, with ~75% of animals becoming long-term survivors following surgical resection (p < 0.05, Fig. 4A). Surviving treated mice were re-challenged with 100,000 4T1 cells in the contralateral mammary fat pads. The majority of IVM + IP-treated mice did not develop new tumors, which is indicative of the development of long-lasting tumor-specific immunity (Fig. 4B), while IP-treated and control naïve nice all developed tumors. Splenocytes from these animals were reactive (via ELISPOT) against 4T1 cells, demonstrating evidence for anti-tumor T cell responses in the IVM + IP-treated animals (Fig. 4C). These results suggest that the IVM + IP combination treatment is effective in the neoadjuvant setting and induces potentially stronger protective anti-tumor immunity in responders. Fig. 4: Combination ivermectin and IP therapy in the neoadjuvant, adjuvant, and metastatic settings. figure 4 A Survival of animals following surgical resection of primary tumor (on day 16 post tumor inoculation). B Induction of protective immunity in treated mice that survived beyond day 80, then re-challenged with 4T1 cells on the contralateral mammary fat pad. C IFNγ ELISPOT analysis of 4T1-reactive splenocytes in treated animals. Mean ± s.d., n = 5 mice, pooled data from two independent experiments; *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001. D In vivo bioluminescence imaging of mice (on day 17 post surgery and after completion of the entire treatment schedule) treated with ivermectin, anti-PD1, ivermectin + anti-PD1 ± IL-2 (IP), or control in the adjuvant setting. Mean ± s.d., n = 5 mice, pooled data from two independent experiments. E Survival of animals in the adjuvant setting following surgical resection of primary tumor burden and treated starting 2 days after with ivermectin, anti-PD1, ivermectin + anti-PD1 ± IL-2 (IP), or control; n = 5 mice per group, two-tailed log-rank test; **p ≤ 0.01, ****p ≤ 0.0001. F In vivo bioluminescence imaging (on day 17 post surgery and after completion of the entire treatment schedule) of mice with documented metastasis, then treated with ivermectin, anti-PD1, ivermectin + anti-PD1 ± IL-2 (IP), or control. Mean ± s.d., n = 5 mice, pooled data from two independent experiments. G Kaplan–Meier survival analysis of mice in the metastatic setting treated with ivermectin, anti-PD1, ivermectin + anti-PD1 ± IL-2 (IP), or control; n = 5 mice per group, two-tailed log-rank test; *p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001. Full size image Surgery remains the primary treatment for breast cancer; however, relapse is common necessitating adjuvant therapy in high-risk patients post-surgery. We assessed the efficacy of ivermectin, anti-PD1, and recombinant IL-2 alone or in combination as adjuvant immunotherapy after surgery. 4T1 cells expressing luciferase (0.5 × 106, 4T1-Luc) were injected into the mammary pad of female BALB/c mice and allowed to grow into palpable tumors over 10 days, after which tumors were surgically resected. Treatment was initiated on day 2 following surgery to mimic adjuvant therapy (schema in Fig. S1A). Development of recurrence and metastasis was monitored at multiple time points via bioluminescence imaging (day 17 shown, Fig. 4D), then animals were monitored until they met euthanasia criteria based on decline in body condition score and signs of morbidity. Treatment with anti-PD1 or IVM alone led to similar survival as untreated controls (Fig. 4E). Animals treated with the combination of ivermectin and anti-PD1 (with or without IL-2) had significantly prolonged survival, with ~40% of animals becoming long-term survivors (p < 0.001, Fig. 4E). Through statistical modeling, the ivermectin and anti-PD1 combination was found to be highly synergistic compared to IVM or anti-PD-1 alone (submodel p = 0.007, FDR 2%, Table 2). Interestingly, addition of IL-2 did not further enhance the survival benefit from the ivermectin and anti-PD1 combination (submodel p = 0.51, FDR 67%, Table 2). These data demonstrate that treatment with ivermectin and anti-PD1 (with or without IL-2) is effective in the adjuvant setting, without evidence for drug-related or synergistic toxicity based on parallel body weight observations (Fig. S1B). Table 2 Relapse-free survival in adjuvant setting. Full size table Metastasis is the main cause of death in cancer patients including breast cancer. To test the efficacy of this combination in the metastatic setting, we delayed treatment until at least 25% of animals post-surgery had detectable metastasis (generally day 7 after surgical resection of primary tumor). Progression of metastasis was monitored via bioluminescence imaging (schema in Fig. S1A), and animals were monitored until they met euthanasia criteria based on decline in body condition score and signs of morbidity (examples shown in Fig. 4F). All untreated animals required euthanasia due to metastatic disease around day 20–40 following surgical resection of primary tumor (Fig. 4G). Treatment with IVM alone led to modest, non-significant prolongation of survival as compared to untreated controls (Fig. 4G). Survival was slightly prolonged in animals treated with anti-PD1 only (p < 0.05), but all animals required euthanasia by Day 60 as in the IVM alone group. Survival was significantly prolonged in animals treated with ivermectin and anti-PD1 (p < 0.001), or ivermectin, anti-PD1, and IL-2 (p < 0.01) as compared to untreated controls (Fig. 4G). Approximately 40% of animals on the combination therapy become long-term survivors. The combined effect of IVM and anti-PD-1 on survival in the metastatic setting was again found to be highly synergistic compared to IVM or anti-PD-1 alone (submodel p < 0.001, FDR < 1%, Table 3). As in the adjuvant setting, addition of IL-2 did not further enhance the survival benefit from the ivermectin and anti-PD1 combination (submodel p = 0.64, FDR 73%, Table 3). These data demonstrate that treatment with ivermectin and anti-PD1 (with or without IL-2) is also effective in the metastatic setting. Table 3 Relapse-free survival in metastatic setting. Full size table Discussion Since its discovery in the mid-1970s, ivermectin has been used safely by over 700 million people worldwide to treat river blindness and lymphatic filariasis26; it is inexpensive and accessible. Our results demonstrate that treatment with ivermectin induces robust T cell infiltration into breast tumors via induction of ICD, thus turning “cold” tumors “hot”. Unlike conventional chemotherapy drugs, this agent has the added benefit of not suppressing host immune function, but rather has beneficial immunomodulatory effects—making it a promising and mechanistic partner for immune checkpoint blockade. The release and accumulation of high levels of extracellular ATP has emerged as a key characteristic feature of the tumor microenvironment27, and a hallmark of ICD. We and others have previously shown that ivermectin is a positive allosteric modulator of purinergic signaling and the ATP/P2X4/P2X7/Pannexin-1 axis which operates in both cancer and immune cells20,28. In murine splenocytes treated ex vivo, we showed that ivermectin can selectively target various immune subpopulations in a P2X7-dependent fashion (Fig. 2B, C) and has immune-potentiating activities associated with augmented ratios of immune effectors versus immunosuppressive populations, including Tregs and myeloid cells (Fig. 2A, D). The observed selective targeting of different immune populations by ivermectin is consistent with previous reports demonstrating that mouse splenic Tregs (CD4+CD25+) have higher sensitivity to increasing (>100 μM) doses of extracellular ATP compared to CD8+ and CD4+CD25− T cells29. This differential sensitivity to extracellular ATP is P2X7-dependent and directly associated with levels of surface P2X7 receptor expression (CD4+CD25+ > CD4+CD25− > CD8+ T cells). Recent reports showed that the ATP/P2X7 axis also operates in MDSC and MDSC-mediated immunosuppression30,31. This is consistent with our finding that ivermectin can selectively target expanded myeloid cells isolated from tumor-bearing mice ex vivo in a P2X7-dependent fashion. Further research will be needed to elucidate the relative sensitivities of different subsets of MDSC and tumor-associated macrophages/neutrophils (TAMs/TANs) to ivermectin, as well as to validate the in vivo effects of ivermectin on various myeloid subsets within the tumor microenvironment and systemically. While differential ATP/P2X7-dependent cytotoxicity may be one possible explanation for the immunomodulatory effects of ivermectin in vivo, recent reports also implicate ATP release and P2X4-dependent signaling in the CXCL12/CXCR4-mediated migration and inflammation-driven recruitment of T cells32. The role of P2X4 in T cell activation, proliferation, and migration was particularly pronounced in CD4 T cells, which is consistent with our own data demonstrating ivermectin to be particularly potent at increasing the CD4+ Teff/Treg ratios in ex vivo treated splenocytes (Fig. 2D) and augmenting intra-tumoral infiltration with CD4+ T cells (Fig. 3D). Thus, infiltration of tumors by T cells in ivermectin-treated mice may reflect a combination of selective depletion of suppressive cells as well as recruitment effects. The synergistic activity between ivermectin and anti-PD-1 checkpoint blockade at driving T cell infiltration into the tumor microenvironment is particularly intriguing as PD-1 functions as a negative feedback regulator of TCR signaling. P2X4/P2X7-gated Pannexin-1 (PANX1) opening and ATP release play a central role in T cell activation by providing a feed-forward loop for TCR-initiated and ATP-driven ATP release at the immunological synapse. The ability of ivermectin as an allosteric modulator of P2X4/P2X7/PANX1 receptors to modulate purinergic signaling operating in both cancer and immune cells therefore may be enhanced by elevated levels of ATP within the tumor microenvironment and the immunological context, including magnitude of chemokine/TCR signaling and chemokine/TCR-driven ATP release. Consistent with the latter possibility, we demonstrated that the potentiating effect of ivermectin on the Teff/Treg ratio appears to be stronger and sustained in the presence of T cell stimulation via PHA (Fig. 2D). Further studies will be needed to unravel how these multi-faceted effects of ivermectin to induce immunogenic cancer cell death, differentially modulate immune cells, and harness the ATP-rich tumor microenvironment may all contribute to its ability to synergize with immune checkpoint blockade in vivo. Immune checkpoint inhibitors (ICIs) are effective as single agents only in a small subset of cancer patients. Hundreds of clinical trials are currently testing various combinations of ICIs with FDA-approved or experimental agents. Such combinations are mainly put together based on partial efficacy of the partnering agent with little or no mechanistic rationale for synergy. Importantly, recent analyses found no evidence from any trial data reported to date that ICIs are synergistic or additive with other drugs33, but instead synergistic toxicity may be observed34,35. We showed that ivermectin represents a rational mechanistic partner for immune checkpoint blockade, demonstrating bona fide synergy when neither agent worked alone. Synergy between PD-1 blockade and ivermectin is mechanistically associated with the ability of the ivermectin to drive immunogenic cancer cell death and T cell infiltration into tumors, thus converting “cold” tumors “hot”36. In contrast, evaluation of the therapeutic potential of PD-1 blockade combined with established first line standard of care chemotherapeutic agents that are also known to induce ICD, such as doxorubicin, in similar 4T1 models failed to achieve durable responses or complete tumor regressions37,38. Doxorubicin alone did not induce significant T cell infiltration into tumors and showed no synergistic activity with PD-1 blockade. Failure of doxorubicin to turn tumors “hot” or effectively synergize with PD-1 blockade likely reflects its direct immune effects. Doxorubicin at concentrations corresponding to plasma levels achieved in patients39 causes significant cytotoxicity to peripheral blood mononuclear cells (PBMCs) (Fig. S3), while physiologically relevant levels of ivermectin showed no significant effects. As demonstrated in our current study, the combination of ivermectin and PD-1 checkpoint blockade led to complete regression of the primary tumor in a significant fraction of animals, and with protective anti-tumor immunity in the responders. We went on to demonstrate that this novel combination is effective in the neoadjuvant, adjuvant, and metastatic settings that mimic clinical situations in which it may be used. Based on its novel dual mechanisms of action in cancer, ivermectin may also potentiate the anti-tumor activity of other FDA-approved ICIs. Lastly, ivermectin is inexpensive, making it attainable for everyone including cancer patients in developing countries. The preclinical findings we present suggest that the combination of ivermectin and anti-PD1 antibody merits clinical testing in breast cancer patients. Methods Mice and treatment Female BALB/c mice were purchased from Charles River Laboratories at 5–8 weeks of age and housed in City of Hope’s animal care facilities under pathogen-free conditions. All procedures were performed under approval from City of Hope’s Animal Care and Use Committee. Mice were inoculated with 100,000 4T1 breast cancer cells in the right mammary fat pad, then palpated to check for tumor engraftment before commencing their assigned treatment regimen. Treatments included: 5 mg/kg of ivermectin (Sigma Aldrich, St. Louis MO) given via oral gavage daily for 6 days; 10 mg/kg of anti-PD1 (BioXCell, West Lebanon NH) treatment given subcutaneously once weekly; MSA-IL2 administered at 1.5 mg/kg by intraperitoneal injection in 50 µL of sterile PBS once weekly; combination treatments; or no treatment (Fig. S1). Ivermectin was solubilized in 45% (2-Hydroxypropyl)-β-cyclodextrin (Sigma Aldrich, 332593-1KG) and was used at half the daily dose of 10 mg/kg, as previously described40. The dose and frequency of ivermectin treatment was optimized for maximum efficacy and absence of visible drug-related toxicity based on animal observations and body weight measurements. Tumor growth was measured 2–3 times a week with a digital caliper for up to 56 days. Mice were euthanized when tumor growth reached 1.5 cm in length or width. Tumor volume was calculated as (length × width2)/2. Metastasis experiments were performed by injecting 0.5 × 106 luciferase expressing 4T1 tumor cells (4T1-Luc) subcutaneously in the mammary gland of female BALB/c mice, followed by surgically resection of the primary tumor on day 14 after inoculation. In vivo bioluminescence imaging was used to monitor metastatic outgrowth, which was carried out on a Lago X optical imaging system (Spectral Instruments Imaging, Tucson, AZ). Overall tumor burden per mouse was assessed weekly via bioluminescence imaging. Recurrence of primary tumor was recognized when the animal’s luciferase value exceeded 600,000 photons/s/cm2/steradian, a threshold chosen because it was well above the lower limit of reproducible detection (510,000) and because, in optimization experiments, 600,000 was the lowest threshold consistently followed by ever-increasing values and eventually death. There was no significant toxicity following treatment with oral ivermectin combined with systemic anti-PD1 and IL-2 as measured by weight loss (Fig. S1). Neoadjuvant setting mice and treatment Female BALB/c mice were purchased from The Jackson Laboratories at 6–8 weeks of age and maintained in animal care facilities under pathogen-free conditions at the Massachusetts Institute of Technology. All procedures were performed under approval from MIT’s Animal Care and Use Committee. An inoculum of 0.5 × 106 4T1-Luc tumor cells were injected subcutaneously (s.c.) in the mammary gland in 100 µL of sterile PBS. Tumor onset was monitored by palpation (usually 3–5 days after inoculation). Six days following inoculation, mice were randomized into treatment groups and treatment was performed as indicated in Supplementary Fig. 2. A dose of 5 mg/kg ivermectin was administered by oral gavage in 50 µL of sterile PBS. Anti-PD-1 (clone RMP1-14, BioXCell) was administered at 10 mg/kg by intraperitoneal injection in 50 µL of sterile PBS. MSA-IL2 was administered at 1.5 mg/kg by intraperitoneal injection in 50 µL of sterile PBS. Surgical resection of primary tumor was performed on day 16 following tumor inoculation. Mice were injected with the analgesic sustained-release Buprenorphine (ZooPharm, 1 mg/kg body weight) and meloxicam (2 mg/kg body weight) by subcutaneous injection. Animals were anesthetized with isoflurane and complete anesthetization was confirmed by lack of a toe pinch reflex. The surgical area was shaved and sterilized by swabbing with alternating application of betadine surgical scrub and 70% ethanol. The tumor and surrounding mammary fat pad was removed by blunt dissection using autoclaved surgical instruments (Braintree Scientific). Wounds were closed using 4-0 nylon monofilament sutures with a 3/8 reverse cutting needle (Ethilon). Mice were monitored for consciousness in a warm, dry area immediately post-operation. Thereafter, mice were dosed with meloxicam (2 mg/kg body weight) at 24 h intervals for 3 days post-surgery. Sutures were removed at 7–10 days post-operation. Mice were monitored for development of metastasis starting at day 10–14 following surgical resection of the primary tumor. Animals were injected i.p. with sterile-filtered D-luciferin (Xenogen) in PBS (150 mg/kg body weight in 200 µL) and anesthetized with isoflurane. Bioluminescence images were collected at 10 min following injection with a IVIS Spectrum Imaging System (Xenogen). Acquisition times ranged 10–30 s. Images were analyzed using Living Image software (Xenogen). Animals were monitored daily for morbidity and euthanized if signs of distress were observed, including but not limited to difficulty in ambulating or breathing, significant weight loss (>20% starting body weight), poor body condition (score <2), or veterinary staff recommendation. Necropsy was performed to confirm presence of visible metastatic nodules. To evaluate response to re-challenge, mice that survived metastasis development following surgical resection or naïve control mice were challenged with a subcutaneous injection of 0.1 × 106 4T1-Luc cells in 100 µL of sterile PBS in the flank opposite the site of the primary tumor. The mice were subsequently monitored every 2–3 days for tumor growth at the inoculation site. ELISPOT assay Target 4T1-Luc cells were treated with mouse IFN-gamma (Peprotech) for 18 h, washed, and irradiated (120 Gy). Splenocytes were isolated from untreated or treated mice on day 16 following to tumor re-challenge. Quantification of IFNγ response was determined using a BD mouse IFNγ ELISPOT kit. Target cells were seeded at 0.025 × 106 cells per well. Effector cells were seeded 1.0 × 106 cells per well. Plates were wrapped in foil and incubated at 37 °C for 24 h and developed following the manufacture’s protocol. Plates were scanned using a CTL-ImmunoSpot plate reader and spots were enumerated using CTL ImmunoSpot software. Tissue staining and quantification Tumors were isolated from mice and sectioned into 5 µm sections for staining with the desired markers (below) using Tyramide Signal Amplification (PerkinElmer, Waltham MA) per manufacturer’s protocol. Whole tumor images were scanned using the Vectra 3 Automated imaging system (PerkinElmer) and quantified using the ImagePro analysis software. Flow cytometry Cell surface markers were stained for 30 min in the dark at 4 °C. Intracellular cytokine staining was performed using the ebioscience Foxp3 staining kit (ThermoFisher Scientific, Waltham MA) per manufacturer’s protocol. The following mouse antibodies from BioLegend (San Diego CA) were used: CD4 (GK1.5); CD8 (53-6.7); Tbet (4B10); Gata3 (16E10A23); Foxp3 (MF-14); IFNγ (XMG1.2); IL-10 (JES5-16E3); IL17 (TC11-18H10.1); and TGFβ (TW7-16B4). RORγt (AFKJS-9) was ordered from eBioscience (ThermoFisher Scientific). To show T cell reactivity, splenocytes were isolated from tumor bearing mice and cultured with 4T1 cells at a ratio of 5:1 (splenocytes to tumor cells) in the presence anti-CD107A/CD107B (ThermoFisher Scientific) and Monensin for 4 h. After 4 h, cells were stained for surface and intracellular markers described above. Flow cytometry analysis of T cell markers on human PBMCs was performed using the following clones: CD8 (RPA-T8); CD4 (SK3); Tbet (4B10); Ki67 (Ki67) from BioLegend; RORγt and granzyme B (GB11) from ThermoFisher Scientific. Statistical analysis Mean values were compared using t tests. Data on tumor volume over time were log-transformed prior to statistical modeling; prior to transformation, values of zero were replaced with 0.1. Complete response (CR) to treatment was defined as permanent shrinkage of tumor volume to zero at any time during follow-up; no tumor that shrank to zero resumed growth. The competing survival outcome was progression, defined as tumor growth beyond 150 mm3, after which tumors never underwent CR but instead became necrotic or large, necessitating euthanasia. The follow-up of subjects that experienced neither CR nor progression was censored at last observation, except when the last available tumor measurement fell just short of the 150 mm3 threshold for progression; in such cases (n = 2, volume 139 and 141 mm3, respectively, at final measurement on day 25), progression was assumed to occur by what would have been the next scheduled measurement. Cumulative incidence of competing outcomes was calculated and plotted according to Gray41. The related outcomes of tumor volume, CR, and progression were modeled jointly42. The submodel of longitudinal tumor volume used linear mixed regression, while the survival submodels of CR and progression used parametric hazard regression with Weibull function. Statistical significance was defined as p < 0.05. A greater-than-additive (synergy) effect of combination therapy was demonstrated when the sum of effects of each drug alone fell outside the 95% confidence interval around the effect of combination therapy. To maximize statistical power and obtain unbiased results despite the non-random missingness of longitudinal data due to death, each pair of outcome measures per trial was modeled jointly42. Each joint model included a linear mixed effects submodel of the longitudinal outcome and a survival submodel. 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Article Google Scholar Hochberg, Y. A Sharper Bonferroni procedure for multiple tests of significance. Biometrika 75, 800–802 (1988). Article Google Scholar Draganov, D. Metadata record for the manuscript: ivermectin converts cold tumors hot and synergies with immune checkpoint blockade for treatment of breast cancer. figshare https://doi.org/10.6084/m9.figshare.13568018 (2021). Small heat shock proteins (sHSPs) are ATP-independent molecular chaperones characterized by a conserved α-crystallin domain located between a highly flexible, variable C-terminal region and a poorly conserved, disorganized N-terminal domain (NTD) (1). Like other chaperones, sHSPs bind proteins to facilitate substrate refolding, serving as ubiquitous components of protein quality control networks. HSP27 (also referred to as HSPB1) assembles into oligomers that interact with misfolded or specific client proteins to regulate proteostasis and cytoskeleton organization, as well as prosurvival signaling and transcriptional pathway activation (2). Assembly and depolymerization of HSP27 subunits are dynamic and context-dependent processes, mediated by phosphorylation of 3 serine residues (Ser15, Ser78, Ser82) in its NTD by kinases (e.g., MAPKAP2, p38 MAPK, p90RSK) activated by varied stress stimuli (e.g., hypoxia, heat shock, nutrient deprivation, anticancer treatments) (3). High levels of HSP27 are linked to neurodegenerative (Alzheimer, Parkinson) and other aggregopathies (e.g., cystic fibrosis) as well as inflammatory diseases (4, 5). HSP27 also regulates many hallmarks of cancer and is associated with poor outcome and treatment resistance. Notably, HSP27 phospho-activation facilitates proteostasis, prosurvival, and antiapoptotic pathways, as well as specific oncoprotein (e.g., BRAF, MET, AR, EGFR) interactions (6, 7). In many model systems, HSP27 suppression sensitizes cancer cells to therapy (8). Collectively, these broad effects of HSP27 on cancer cell proliferation and survival position HSP27 as a potential therapeutic target. Since phospho-activation of HSP27 modulates its dynamic oligomer assembly and defines its affinity to client proteins (9), targeted inhibition of HSP27 phosphorylation would suppress many oncogenic pathways that facilitate treatment resistance. However, because HSP27 is intrinsically disordered and lacks an ATP-binding domain (1), rational discovery of small-molecule inhibitors of HSP27 poses significant challenges. Apatorsen (OGX-427) is a second-generation antisense oligonucleotide targeting HSP27 mRNA with anticancer activity in many preclinical models (7, 10, 11). Randomized phase II trials in castration-resistant prostate cancer reported higher response rates with apatorsen plus prednisone compared with prednisone alone, consistent with inhibition of HSP27 shuttling of androgen receptor (AR) into the nucleus (7, 12). Another phase II trial randomized 200 patients with advanced urothelial carcinoma to docetaxel plus apatorsen versus docetaxel alone and reported survival benefit for patients receiving apatorsen (13). While these studies report the first signals of activity for an HSP27 inhibitor in cancer, antisense drugs have variable intratumor distribution, and a more potent and convenient orally available small molecule would be preferable. To identify small-molecule inhibitors of HSP27, we followed a course of drug screening coupled with structural modeling approaches to resolve a targetable phosphorylation pocket in its NTD, identify ivermectin (IVM) as an inhibitor of HSP27 phospho-activation, and delineate its biochemical and functional mechanisms of action in cancer. Results Screening for HSP27 small-molecule inhibitors Combined screening using fluorescence resonance energy transfer (FRET) and fluorescence polarization (FP) assays was initially used to identify drugs in the Prestwick Chemical Library (Prestwick Chemical) that inhibit HSP27 functions (Supplemental Figure 1A; supplemental material available online with this article; https://doi.org/10.1172/JCI130819DS1); compounds passing both FRET and FP screening criteria were further analyzed in secondary assays (described below). Of 1280 Prestwick molecules evaluated, 6 passed initial FRET and FP screening (Supplemental Table 1), but only IVM passed confirmatory secondary assays, and was subsequently found to functionally inhibit HSP27. IVM is a member of the avermectin family of macrocyclic lactones (herein called IVM analogs) that similarly inhibit HSP27 in FRET and FP assays (Supplemental Table 2). Since rapid exchange of HSP27 subunits is essential for HSP27 chaperone activity (14), FRET, which relies on the distance-dependent transfer of energy between donor and acceptor molecule, was used to investigate drug-HSP27 interaction and effect on subunit exchange kinetics. IVM interfered with HSP27 fluorescence quenching when pools of Rhodamine Red–labeled HSP27 (HSP27-RR, electron donor) and QSY21-labeled HSP27 (HSP27-QSY21, electron acceptor) were mixed (Figure 1A). IVM also dose-dependently inhibited HSP27 interaction with unfolded insulin (Figure 1B), an established HSP27-client protein used to study HSP27 inhibition of misfolded protein precipitation, as shown by the FP assay (1, 15). Figure 1. Ivermectin binds to HSP27 phosphorylation pocket. Figure 1 Open in a new tab (A) FRET assay using HSP27 labeled with QSY21 as negative control (blue triangles), and HSP27-RR alone (green triangles) or combined with IVM (brown circles) as positive controls. Red squares represent IVM effect on subunit exchange. (B) FITC-insulin precipitation in the presence of DMSO or increasing concentrations of IVM. (C) BLI dose-response curves reflect direct binding of IVM to purified HSP27 protein. (D) Protein aggregation in HSF1–/– MEF cells incubated with HSP27 (left) or HSP70 and HSP90 (middle and right) in the presence of IVM (gray) or DMSO (red). Protein lysate without chaperones was used as negative control (black). Results were normalized to HSP values alone. For HSP90 the effect of IVM was compared with that of 17AAG (blue). (E) Ribbon drawing of the HSP27 24-mer down its 3-fold symmetry axis. Arrows represent bisecting 2-fold axes. Monomers A are shown in green and monomers B in brown. (F) Magnified view of the interface between monomers in a dimer unit showing the phosphorylation pocket. Spheres indicate S78 and S82, and the mesh represents the WDPF motif of each monomer. Peptides 6 and 11 conferring chaperone activity are shown in blue and red, respectively. (G) Ribbon representation of the NTD of monomer A (green) fitting into a pocket created by its related monomer B and a neighboring monomer B NTD. (H) Left: Predicted binding pocket between the 2 NTDs of an HSP27 dimer: monomer A (gold), monomer B (cyan), and IVM (violet). Right: S78 and S82 and other amino acids involved in IVM interaction. Hydrogen bonds are shown in green. (I) BLI reported binding of IVM to the HSP27 phosphorylation pocket mutants S78A-S82A (left) and S78D-S82D (right). These screening assays identify IVM analogs as possible HSP27-interacting molecules; secondary assays were subsequently used to evaluate inhibitory effects of IVM on HSP27 client–protein interactions. Ivermectin directly binds HSP27 and prevents its interaction with misfolded proteins Direct binding of IVM to HSP27 was confirmed by biolayer interferometry (BLI). HSP27 was expressed and purified with a biotinylated tag and immobilized on streptavidin biosensors. Sensors were then dipped in successive wells containing increasing concentrations of IVM. Binding of IVM to protein changes the spectral pattern of reflected light. BLI indicates dose-response binding of IVM to HSP27 (Figure 1C), while the AR antagonist enzalutamide, used as a negative control, did not bind HSP27 at any concentration (Supplemental Figure 1B). To determine effects of IVM on HSP27 inhibition of protein aggregation, HSF1–/– MEF cells negative for HSP27, HSP70, and HSP90 were used as a source of cellular proteins (16). IVM reduced HSP27-mediated (but not HSP70- or HSP90-mediated) inhibition of protein aggregation in HSF1–/– MEF cell protein extracts (Figure 1D). IVM effects on HSP27-mediated inhibition of protein aggregation were similar to 17AAG suppression of HSP90-mediated inhibition of protein aggregation. Since actin is an established interaction partner of HSP27 (17), an actin-pyrene assay was used to evaluate HSP27-regulated actin polymerization and its functional inhibition by IVM (Supplemental Figure 1C). IVM also reduced trypsin degradation of HSP27 (Supplemental Figure 1D). Collectively, these data demonstrate that IVM inhibits HSP27 interaction with specific client proteins. Determination of oligomeric structure of HSP27 To facilitate study of HSP27 function and specifics of its inhibition by IVM, x-ray crystallography was used to resolve HSP27 structure. These studies define an oligomeric complex of HSP27 formed by 24 monomers in a spherical configuration (Figure 1E). Polypeptide chains form a layer of approximately 40 Å thickness about the surface of this spheroid, which has a central void of approximately 67 Å diameter. Notably, this complex is composed of 12 dimers wherein the interactions between 2 identical monomers (A and B) are distinct from the interactions formed with other dimer units. Overall the spheroid formed has a central 3-fold axis bisected by perpendicular 2-fold axes. Four open surface channels of approximately 28 Å diameter each are found about the surface of this complex and could potentially allow for access by small molecules. The HSP27 monomer. The HSP27 protein is composed of 205 amino acids constituent in a single polypeptide chain. At its structural core (residues 94–168) there is a beta-sandwich immunoglobulin-like domain that bears a close relationship to the structure and primary sequence of α-crystallin (99% similarity) (18). Emanating from this core and projected to the outer surface are the N- and C-terminal regions, consisting of residues 1–93 and 169–205, respectively. The α-crystallin domain of HSP27 is well resolved in our structural determination, whereas the remaining polypeptide chain demonstrates greater mobility. Residues found to be particularly mobile are the 55–93 and 190–205 positions. Some loop segments of the HSP27 protein were not built into our model because of positional disorder in those regions, and an average of 39 residues per chain were missing. Dimer relationships. Within the overall 24-meric structure of HSP27, its dimer units are involved in 2 types of interactions. Within the dimer itself there is a 2-fold relationship between identical monomers that leads to very compact structural packing. It is at this interface that much of the functionality of HSP27 would seem to reside. Notably, S78 and S82 in each monomer of HSP27 are positioned in a common surface cavity, suitable for small-molecule binding (Figure 1F). An increased degree of phosphorylation at these serine residues as well as at S15 drives dissociation of oligomeric HSP27 (9). Conversely, oligomeric complex formation is favored by a lack of phosphorylation at these residues (19). Coincident with oligomeric dissociation is the exposure of the hydrophobic surfaces of 2 nearby peptide strands composed of residues 42–56 (peptide 6) and residues 87–102 (peptide 11), which contributes to enhanced HSP27 chaperone activity with insulin and actin (17) (Figure 1B and Supplemental Figure 1C). Modeling studies also established that the floor of the intersubunit cavity within HSP27 dimers is flanked on either side by the corresponding WDPF motif (residues 16–19) of both monomers and its associated S15 phosphorylation site. Our modeling does not reveal a direct contribution of S15 to the cavity formation (Figure 1F). The N-terminal extension of this polypeptide chain region is of particular relevance, as it forms interactions with not only the adjacent monomer, but also nearby dimer units (Figure 1G), and therefore appears to form a significant part in dimer-dimer association and oligomerization interactions. The presence of the WDPF motif along with the phosphorylation state of above-mentioned serine residues has also been reported to be a key factor in HSP27 oligomerization (20). Placement of ivermectin binding between the NTDs of an HSP27 dimer Our structural studies support the hypothesis that chemicals binding the phosphorylation pocket within HSP27 dimers comprising S78 and S82 of each monomer would inhibit phosphorylation and oligomer dissociation events and therefore block HSP27 function. Docking studies used to define specific interaction of IVM in this pocket indicate favorable hydrophobic interactions with H90, V85, I63, and V68 of monomer A and formation of hydrogen bonds with G54, S78, and Q190 residues of monomer B (Figure 1H). This binding model predicts that IVM would block phosphorylation activity about S78 and S82. Further evidence of the propensity for IVM to bind at this site is apparent from 3 independent molecular dynamic (MD) simulations (Supplemental Figure 1E and Supplemental Videos 1 and 2) of the HSP27 24-mer in complex with 12 docked IVM molecules over 300 nanoseconds. These computations indicate that during each trial IVM contacted at least 1 of the 4 serine residues (Figure 1H and Supplemental Figure 1F). Direct binding of IVM to the phosphorylation pocket of HSP27 was confirmed by BLI experiments using HSP27 double mutants S78D-S82D (chemically mimicking phosphorylated HSP27 and stabilizing HSP27 as dimers) and S78A-S82A (unphosphorylatable HSP27, stabilizing HSP27 as a 24-mer). These HSP27 double mutants were expressed and purified with a biotinylated tag and then exposed to increasing concentrations of IVM. Binding of IVM to S78A-S82A unphosphorylatable mutant was similar to its binding to wild-type (WT) protein (Figure 1I, left); in contrast, IVM binding affinity was significantly lower with the phospho-mimic double mutant S78D-S82D (Figure 1I), indicating that IVM binds to HSP27 in its non-phosphorylated form. Ivermectin inhibits HSP27 phosphorylation and depolymerization Consistent with predicted computational modeling described above, IVM potently inhibited HSP27 phosphorylation in lung and prostate cancer (Figure 2A) as well as bladder and breast cancer (Supplemental Figure 2A) cells. Notably, IVM affected mainly phosphorylated HSP27 (p-HSP27), while total HSP27 protein levels were marginally affected. Of the avermectins, IVM most potently suppressed p-HSP27 levels (Supplemental Figure 2B). Since IVM similarly inhibits phosphorylation of S78 and S82, while levels of S15 were only marginally altered, the effect of drug on S78 was selected as a readout in subsequent experiments. Figure 2. Ivermectin inhibits phosphorylation and oligomerization of HSP27 in cancer cells. Figure 2 Open in a new tab (A) Protein expression of phosphorylated (S78) and total HSP27 was evaluated by Western blotting after heat shock (42°C for 1 hour) in lung HCC-827 and prostate 22RV1 and PC3 cancer cells after treatment with increasing doses of IVM for 24 hours. (B) Quantification of HSP27 phosphorylation expression assessed by Western blotting after in vitro incubation with MAPKAP2 kinase in the presence of DMSO or IVM, at time 0 and after 24 hours. Results were averaged and normalized as a percentage of the control at 24 hours. Data shown as mean ± SEM; *P < 0.05 by unpaired 2-tailed Student’s t test with Welch’s correction; n = 3 independent experiments. RIU, relative intensity units. (C) Expression of HSP27 protein fractions of glycerol gradient from PC3 (top panel) and HCC-827 (bottom panel) cells treated with DMSO (CTRL) or IVM (5 μM for 24 hours). (D) PC3 (top panel) and HCC-827 (bottom panel) cells treated with DMSO (CTRL) or IVM (5 μM for 24 hours) were exposed to heat shock (42°C for 1 hour), fixed using cold methanol, and stained for HSP27 (green). DAPI (blue) was used for nuclear staining. Scale bars: 20 μm. p38 MAPK phospho-activates the serine/threonine MAPKAP2 kinase that directly phosphorylates serine and threonine residues in HSP27 (20). IVM did not alter protein levels of phospho–p38 MAPK (Supplemental Figure 2C), supporting direct suppression of HSP27 phosphorylation rather than suppression via upstream modulation of kinases that phosphorylate HSP27. Indeed, in vitro phosphorylation of purified WT-HSP27 by MAPKAP2 kinase was significantly decreased by IVM (Figure 2B), supporting direct inhibition of HSP27 by blocking of MAPKAP2-mediated phosphorylation. Phosphorylation of HSP27 facilitates disassembly of HSP27 into smaller oligomers and enhances subunit exchange to promote HSP27 chaperone activity (9, 19). Consistent with its effects on phosphorylation, IVM shifted HSP27-oligomer equilibrium in prostate and lung cancer cells, stabilizing HSP27 in larger (less active) complexes (Figure 2C). Ivermectin inhibits HSP27 nuclear translocation Upon heat shock, HSP27 is phosphorylated and traffics to the nucleus to chaperone proteins required for mRNA processing during cellular recovery from stress. Consistent with its inhibitory effects on HSP27 phosphorylation and depolymerization, IVM clearly prevented heat shock–induced nuclear translocation of HSP27 in prostate and lung cancer cells (Figure 2D). Collectively, the above screening, biochemical, structural, and computational studies identify IVM as an inhibitor of the HSP27 NTD, blocking access of MAPKAP2 kinase to S78 and S82 to prevent HSP27 phosphorylation and depolymerization. Additional biochemical studies were performed to define IVM effects on HSP27-mediated cellular and oncogenic pathways. Ivermectin inhibits HSP27-regulated epithelial-mesenchymal transition and proteostasis HSP27 plays roles in both epithelial-mesenchymal transition (EMT) (2) and proteostasis (4). Similarly to HSP27 silencing (2), IVM reduced cell migration and increased E-cadherin expression in prostate and lung cancer cells, both markers of EMT (Supplemental Figure 2, D and E). Like HSP27 silencing (10), IVM also led to endoplasmic reticulum (ER) stress with increased PERK and ATF4 expression (Supplemental Figure 2F). The inhibition of HSP27-regulated EMT and ER stress provides additional evidence of the on-target inhibition of IVM. IVM inhibition of HSP27 is synergistically lethal to cells with oncogenic activation of EGFR or AR IVM inhibits HSP27-regulated EGFR activity. HSP27 is associated with resistance to inhibitors of EGFR and HER2, key oncogenic drivers in lung and breast cancer (11, 21). HSP27 inhibition, by IVM or si-HSP27, reduced EGFR phosphorylation in HCC-827 lung cancer cells harboring constitutionally active mutated EGFR, or after EGF stimulation in WT-EGFR SW48 colorectal cancer cells (Figure 3A). No changes were observed in EGFR mRNA or total protein levels (Supplemental Figure 3A and Figure 3A). Coimmunoprecipitation and proximity ligand assay demonstrate that HSP27 directly interacted with EGFR in HCC-827 cells and this was inhibited by IVM (Figure 3B and Supplemental Figure 3B). EGFR inhibition by erlotinib also decreased HSP27 interaction with EGFR (Figure 3B, bottom), suggesting that HSP27 interacted mostly with phosphorylated EGFR (p-EGFR); inhibition of EGFR phosphorylation, either directly with erlotinib or indirectly using IVM via HSP27, perturbed this interaction. Figure 3. IVM reduces EGFR phosphorylation levels via HSP27 regulation of SHPTP1. Figure 3 Open in a new tab (A) Immunoblotting for p-EGFR and total and p-HSP27 after si-HSP27 (20 nM) or IVM (5 μM) in HCC-827 cells (left) or IVM (2.5 μM) before and after EGF stimulation (50 μg/mL) in SW48 cells (right). DMSO was used as control. (B) Top panels: Coimmunoprecipitation of HSP27 and EGFR with and without IVM (5 μM) for 24 hours in HCC-827 cells. IgGs were used as negative control. Bottom panels: Proximity ligation assay between HSP27 and EGFR in HCC-827 cells. Confocal microscopy was used to detect interaction (red dots). DNA was counterstained with DAPI (blue). Erlotinib was used as a negative control. Scale bars: 20 μm. (C) Left panel: Immunoblotting for SHPTP1 and p-HSP27 in HCC-827 cells as in A. Right panel: Immunoblotting for p-EGFR, SHPTP1, and p-HSP27 in wild-type and HSF1–/– MEF cells transfected with either empty (E) or HSP27 plasmid. (D) SHPTP1 phosphatase activity in HCC-827 cells after DMSO or IVM (5 μM). Data shown as mean ± SEM; ***P < 0.001 by unpaired 2-tailed Student’s t test with Welch’s correction; n = 3 independent experiments. (E) Cell survival of HCC-827 and SW48 cells measured by crystal violet assay after 4 days of treatment with DMSO, IVM, erlotinib, or cetuximab and their combination with IVM. Data shown as mean ± SEM; **P < 0.01, ***P < 0.001, ****P < 0.0001 by ANOVA with Bonferroni’s correction. (F) Immunoblotting for EGFR, p-EGFR, HSP27, p-HSP27, ERK, p-ERK, AKT, p-AKT, PARP, and cleaved PARP in HCC-827 cells treated as in E for 24 hours. (G) Tumor volume of HCC-827 xenografts treated with 15 mg/kg erlotinib, 10 mg/kg IVM, or a combination of both. Data shown as mean ± SEM; *P < 0.05, ****P < 0.0001 by ANOVA with Tukey’s correction; n = 8. (H) Immunohistochemical analysis of p-HSP27, p-EGFR, SHPTP1, and TUNEL staining of HCC-827 xenografts. Scale bars: 10 μm. HSP27 regulates EGFR activation by modulating SHPTP1 phosphatase activity. The activation of EGFR is tightly regulated by specific phosphatases that stop its intracellular signaling by dephosphorylating its tyrosine kinase domain (22). SHPTP1 has been demonstrated to significantly reduce EGFR phosphorylation in human colorectal cancer models (23). Therefore, changes in SHPTP1 levels were analyzed after HSP27 silencing or IVM to define how HSP27 regulates EGFR phosphorylation (Figure 3C, left). Low p-EGFR and high SHPTP1 levels were also detected in HSF1–/– MEF compared with wild-type MEF cells, further supporting an inverse relationship between HSP27 and SHPTP1 levels. HSP27 transfection in HSF1–/– MEF cells rescued these effects on p-EGFR and SHPTP1 (Figure 3C, right). SHPTP1 activity increased significantly in HCC-827 cells after IVM (Figure 3D), providing further evidence for HSP27-specific inhibition by IVM, as well as HSP27-mediated stabilization of p-EGFR levels via inhibition of SHPTP1. Similarly to apatorsen (11), IVM augmented inhibitory effects of erlotinib and cetuximab on EGFR signaling, synergistically increasing activity in lung and colorectal cancer cells (Figure 3E); this synergistic effect was also confirmed in other EGFR/HER2-driven tumors (Supplemental Figure 3C). IVM-mediated inhibition of HSP27 phospho-activation enhanced erlotinib-mediated decreases in p-EGFR levels, suppressed AKT and ERK downstream of EGFR, and increased PARP cleavage (Figure 3F). The pharmacokinetic profile of IVM was determined to support tumor growth inhibition studies in vivo. Following oral administration of 10 mg/kg, IVM serum concentrations peaked at 2 hours and remained at 2.13 μg/mL at 24 hours. After 6 days of oral daily treatment, serum concentration was 2.7 μg/mL, and calculated half-life was 26.30 hours. After 6 weeks of 10 mg/kg orally once daily, IVM intratumor concentration in PC3 xenografts was 1.93 μg/g with a serum concentration of 2.85 μg/mL, demonstrating excellent penetration of drug in tumor tissue (Supplemental Figure 3D). To evaluate effects of IVM on progression of constitutionally active EGFRmut lung cancer in vivo, mice bearing HCC-827 xenografts were randomized to vehicle, erlotinib, IVM, or combined erlotinib plus IVM. IVM was well tolerated with no apparent changes in activity and stable body weight of the mice treated with IVM alone or combined with erlotinib, in comparison with vehicle controls (Supplemental Figure 3D). IVM monotherapy reduced HCC-827 tumor p-HSP27 levels and inhibited tumor growth. More importantly, IVM most significantly inhibited tumor growth when combined with erlotinib (Figure 3G), increasing TUNEL staining, reducing p-EGFR staining, and increasing SHPTP1 staining. Although there was a trend toward reduction of Ki-67, this did not reach statistical significance (Figure 3H and Supplemental Figure 3E). p-EGFR staining was also reduced in A549 tumors treated with apatorsen plus erlotinib (11), further supporting the notion that HSP27 inhibition decreases p-EGFR levels in particular when combined with erlotinib (Supplemental Figure 3F). Ivermectin inhibits AR pathway in prostate cancer models. Resistance to AR pathway inhibitors in prostate cancer is associated with AR amplification, mutations, and expression of truncated AR variants, the latter characterized by loss of AR’s ligand-binding domain and constitutively active transcription (24). HSP27 has an established role in AR trafficking and stability, and its inhibition by either apatorsen or si-HSP27 reduces AR protein levels and activity, thereby delaying progression of castration-resistant prostate cancer (CRPC) (7, 8, 25). Similarly to apatorsen (7), IVM significantly reduced AR and prostate-specific antigen (PSA) protein expression in castration-sensitive LNCaP cells and ARF876L protein expression in enzalutamide-resistant M49F cells, an effect that was enhanced in combination with androgen deprivation therapy (in LNCaP cells) or enzalutamide (in M49F cells) (Figure 4A). IVM increased sensitivity of LNCaP cells to androgen deprivation therapy and of enzalutamide-resistant ARF876L M49F cells to enzalutamide (Figure 4B). Interestingly, IVM also decreased AR variant 7 (AR-V7) protein levels in 22RV1 cells, similarly to either si-HSP27 or apatorsen (Figure 4C). The reduction of AR or AR-V7 protein expression was not related to modulation of mRNA levels (Supplemental Figure 4A). The functional effect of IVM on AR-V7 transcriptional activity was also assessed using a PC3V7_3TKNLuc system that incorporates a doxycycline-inducible AR-V7 and probasin-based ARR3tk-Nanoluciferase reporter construct. As shown in Figure 4D, IVM significantly inhibited AR-V7 transcriptional activity compared with PC3_3TKNLuc control. AR-V7 nuclear translocation in 22RV1 cells was also inhibited by IVM, decreasing nuclear AR-V7 levels, with a corresponding increase in the cytoplasmic fraction (Figure 4E). Figure 4. Ivermectin inhibits AR and AR-V7 to potentiate AR pathway inhibitors in vivo. Figure 4 Open in a new tab (A) Immunoblot for p-HSP27, AR, and PSA in LNCaP (left) and enzalutamide-resistant M49F (right) cells after treatment with DMSO, enzalutamide (ENZA) (10 μM), IVM (5 μM), or both for 24 hours. (B) Left panel: LNCaP cell survival in charcoal-stripped serum with or without IVM. Right panel: M49F cells treated with enzalutamide, IVM, or the combination of both for 4 days. Data shown as mean ± SEM; **P < 0.01,****P < 0.0001 by 1-way ANOVA with Bonferroni’s correction. ARPI, androgen receptor pathway inhibitors. (C) AR variant 7 (AR-V7) and full-length (AR-FL) protein expression was analyzed in 22RV1 cells after treatment with DMSO, IVM, si-HSP27, or apatorsen (OGX-427). Scrambled sequences were used as control (si-SCR and SCR-B). Levels of p-HSP27 were used to confirm on-target effect. (D) Dose-dependent inhibition of AR-V7 transcriptional activity measured by luminescence in PC3V7_3TKNLuc cells compared with PC3_3TKNLuc empty control, normalized to DMSO vehicle in each group. n = 3 independent experiments. (E) HSP27 and AR-V7 protein expression was evaluated in cytoplasmic and nuclear fractions of 22RV1 cells treated with DMSO or IVM (2.5 μM) for 24 hours with or without heat shock (42°C for 1 hour). (F) Top panels: Tumor volume and serum PSA quantification of LNCaP xenografts after castration treated with vehicle (CX) or IVM (10 mg/kg). PSA measurement was assessed weekly by ELISA. Lower panels: Western blotting with relative protein expression quantification for AR-FL and AR-V7 of 22RV1 xenografts treated with vehicle (CTRL) or IVM (10 mg/kg). Data shown as mean ± SEM; *P < 0.0001. Tumor volume was compared by ANOVA followed by Tukey’s correction, n = 10 and 6–9 per group for LNCaP and 22RV1, respectively. LNCaP and 22RV1 xenograft models were used to determine effects of IVM on CRPC progression in vivo. Male mice bearing LNCaP xenografts (26) were castrated when tumors exceeded 300 mm3 and randomized to vehicle or IVM administered 10 mg/kg orally 3 times per week. IVM significantly reduced tumor volume and serum PSA levels after castration, delaying time to CRPC (Figure 4F). Moreover, IVM significantly reduced full-length AR (AR-FL) and AR-V7 protein expression and 22RV1 tumor volume growth in vivo (Figure 4F). Collectively these data are consistent with IVM inhibition of HSP27-mediated nuclear transport of AR and its variants to delay prostate cancer progression, defining a novel approach to suppress AR genomic alterations that drive resistance in CRPC. Ivermectin potentiates taxane chemotherapy. HSP27 is linked to resistance to chemotherapy in several cancers (27), and apatorsen plus docetaxel was reported to prolong survival, over docetaxel alone, in a randomized trial of 200 patients with advanced urothelial carcinoma (28). p-HSP27 levels are higher in docetaxel-resistant PC3 cells compared with parental PC3 cells (Figure 5A). IVM sensitized PC3 prostate (Figure 5, B and C) and UC14 urothelial cancer cells (data not shown) to docetaxel and reduced taxane-induced p-HSP27 protein levels. In mice, IVM significantly delayed docetaxel-resistant PC3 tumor progression compared with either drug alone (Figure 5D), lowering p-HSP27 and Ki-67 levels, and increasing TUNEL staining in tumor tissue in the combination arm (Figure 5E and Supplemental Figure 4B). Figure 5. Ivermectin reverses taxane resistance in prostate cancer cells. Figure 5 Open in a new tab (A) Immunoblotting analysis of p-HSP27 and total HSP27 in PC3 parental (WT) and PC3 docetaxel-resistant (DR) cells. (B) Cell survival of PC3-DR treated with docetaxel (DTX), IVM, and the combination of both drugs for 4 days. Data shown as mean ± SEM; ****P < 0.0001 by ANOVA with Bonferroni’s correction. (C) Immunoblotting analysis of p-HSP27 and cleaved PARP in PC3-DR cells treated with increasing doses of IVM for 24 hours. (D) Tumor volume of PC3-DR xenografts treated with control (normal saline), IVM (10 mg/kg orally, every other day), micellar paclitaxel (0.5 mg i.v., 3 times per week, every other week), IVM (10 mg/kg by gavage, 3 times per week), and the combination of both drugs. Data shown as mean ± SEM; ****P < 0.0001; n = 8–14; compared by ANOVA followed by Tukey’s correction. (E) Representative images of immunohistochemical analysis of p-HSP27, TUNEL, and Ki-67 on PC3-DR xenografts treated as specified in D. Scale bars: 100 μm. Discussion Cancer cells co-opt key homeostatic stress responses supporting cell survival and therapy resistance, and molecular chaperones like sHSPs play prominent roles in many stress signaling and transcriptional regulatory networks. In particular, HSP27 is a stress-activated multifunctional chaperone highly expressed in many cancers that regulates misfolded protein and ER stress responses (2, 4), as well as cell survival and oncogenic pathways implicated in cancer progression and treatment resistance (8, 29). HSP27 inhibition, before now only possible by nucleotide-based approaches, induces proteotoxic stress and apoptosis in many varied cancers (27, 30), downregulating oncogenic client proteins and signaling pathways that drive proliferation and EMT (10). Preclinical studies using HSP27 siRNA (25) or antisense-based therapy (10, 11, 27, 30) sensitize cancer cells to combinatorial regimens, and phase II trials of apatorsen in metastatic CRPC and bladder cancer (12, 28) report the first signals of activity for HSP27 inhibitors. While these reports are promising, more potent small-molecule inhibitors are likely to yield improved cancer outcomes. To address challenges of drugging a highly disordered chaperone like HSP27, we integrated structural modeling approaches with biochemical screening of a library of small molecules and specific functional assays. Following initial identification of IVM in FRET and FP screening assays, interaction with and inhibition of HSP27 were confirmed using various complementary biochemical studies. BLI assays of wild-type and mutated forms of HSP27 confirmed direct binding by IVM and contributed to identification of the specific binding region within the NTD. X-ray diffraction data and computer modeling of HSP27 structure defined a spherical 24-mer oligomeric complex with a 3-fold axis and perpendicular 2-fold axis. Each monomer is constructed about a structurally conserved α-crystallin domain exhibiting a 6-stranded beta-sandwich as previously described (31, 32). This conserved domain is flanked by more mobile and variable N- and C-termini that form flexible loops with some sections undefined owing to a lack of electron density as a result of high mobility. The overall structure is similar to an oligomeric lens α-crystallin cryoelectron microscopy structure (18). Each dimer unit in the oligomeric assembly forms a pocket that appears to regulate HSP27 function (Figure 1F). S78 and S82 in the NTD of each monomer are positioned in a common surface pocket, the floor of which is flanked on either side by the corresponding WDPF motif (residues 16–19) of both monomers and its associated S15 phosphorylation site. This portion of polypeptide chain forms interactions with not only the adjacent monomer, but also nearby dimer units, and is likely key to promoting dimer-dimer association and oligomerization. S78 and S82 of the constituent subunits, when phosphorylated, promote dissociation of the HSP27 complex into active-state dimers and the affinity of HSP27 for the spectrum of its context-dependent interactome (9, 15). On the basis of our structural modeling studies, we conclude that IVM binds a pocket comprising S78 and S82 phosphorylation sites between the 2 NTDs of an HSP27 dimer, preventing both phosphorylation and oligomer dissociation events, and therefore HSP27 function. IVM has known anticancer properties, including inhibition of WNT/TCF and HIPPO (33, 34) and induction of mitochondrial ROS and autophagy through degradation of PAK1 (35). Interestingly, HSP27 plays roles in WNT and proteostasis pathways, linking these reported effects to IVM-mediated inhibition of HSP27. While PAK1 can lead to phosphorylation of HSP27 via phospho–p38 MAPK, we established that IVM did not reduce phospho–p38 MAPK levels, indicating that inhibition of HSP27 phosphorylation by IVM is not mediated via PAK1. IVM also inhibits multidrug resistance and efflux of many chemotherapy drugs (36), a mechanism that in part contributes to its activity in docetaxel-resistant PC3 cells. The current work used a broad arsenal of biochemical, structural, and computational methodologies that define a druggable pocket flanked by 2 serine residues in the NTD of HSP27 which, when bound with IVM, inhibits phosphorylation and depolymerization of HSP27 into its active oligomers. IVM inhibits HSP27-specific interaction with client proteins, nuclear translocation, and oncogenic effects similar to those seen with siRNA or antisense drugs. Several chemical compounds (e.g., hardwickiic acid, brivudine) have been reported to inhibit HSP27, but the lack of integrated structural, molecular dynamic, or functional studies fails to confirm specific and direct interaction with HSP27 (37, 38). Given its context-dependent phospho-activation and multiple interactomes, HSP27 can be indirectly affected by compounds through upstream alteration of signaling and/or kinase activity. Thus, xanthones and JCC76 downregulate HSP27 (39, 40), while KRIBB3 reduces HSP27 phosphorylation (41) through unknown mechanisms. Zerumbone and its derivatives inhibit HSP27 through induction of cysteine cross-linking of HSP27 (42), of particular relevance given its 24-mer oligomeric complex (Figure 1E). In the past, computational protocols have been used to build HSP27 monomer models for docking studies of putative HSP27 inhibitors (38, 43). These models, however, have not been published, and could not be compared with the current structural model of HSP27 (38). HSP27 modulates cancer cell sensitivity to targeted drugs by regulating diverse survival pathways. IVM results in phenocopy effects similar to HSP27 silencing (7, 10, 27), inhibiting stress-induced HSP27 phosphorylation and functions in cancer cells, in particular in the dynamic context of oncogenic driver inhibition. IVM inhibited AR and EGFR pathways, and potentiated activity of anti-AR and -EGFR drugs in prostate and lung cancer models, respectively. We also define new molecular crosstalk between HSP27 and selective client-oncoprotein interactions. AR genomic alterations (including amplification, truncated variants, mutations) emerge under selective pressures of castration to drive CRPC progression. IVM, via inhibition of HSP27, reduces AR full-length and AR-V7 protein levels and activity, defining a novel approach to suppress AR variants that drive resistance to AR pathway inhibitors. HSP27 is also highly expressed in, and supports oncogene addiction of, EGFR- and HER2-driven lung and breast cancer (11, 21). Despite initial responses to EGFR inhibitors, resistance-conferring EGFR mutations can emerge and drive progression, which may be overcome by cotargeting of adaptive pathways upregulated by treatment. Similarly to apatorsen (11), IVM synergistically increased activity of erlotinib in lung, cetuximab in colorectal, and trastuzumab in breast cancer cells. We define, for the first time to our knowledge, HSP27-mediated inhibition of the phosphatase SHPTP1, which increases p-EGFR levels and EGFR signaling. IVM inhibits erlotinib-induced increases in HSP27 phospho-activation, which subsequently increases SHPTP1-mediated dephosphorylation of EGFR, synergistically inhibiting tumor growth. In summary, these studies help to define a 3D model of HSP27 with a targetable phosphorylation pocket in its NTD, and identify IVM as an inhibitor blocking access of MAPKAP2 kinase to S78 and S82 to prevent HSP27 phosphorylation, depolymerization, and activation. Inhibition of HSP27 by IVM synergistically enhances AR and EGFR pathway inhibitors, providing a novel repurposing approach to delay emergent resistance to oncogene pathway inhibition. Repurposing IVM in cancer would require higher and longer dosing than its approved use as an antiparasitic. While no significant adverse events were observed in the many in vivo experiments in this study, IVM activates GABA-A receptor activity, which can produce neurologic toxicity. To address these issues, IVM analogs have been designed with lower affinity to the GABA-A receptor while maintaining favorable oral bioavailability, pharmacokinetics, and HSP27 binding affinity, from which a lead compound will be selected for clinical development in combination with AR and EGFR/HER2 inhibitors. Methods Crystallization and x-ray diffraction analyses Purified wild-type HSP27 protein in 20 mM Tris at pH 8.4 was concentrated to 20 mg/mL for crystallization trials. The sitting drop vapor diffusion method was used, with drops containing 1 μL of protein solution, 0.7 μL of 0.5 M ammonium phosphate monobasic, and 0.3 μL of 40% vol/vol tert-butanol. The well solution consisted of 130 μL of 0.5 M ammonium phosphate monobasic. Crystal growth was very slow, with only 2 small crystals (~0.002 mm/side) observed over a 6-month incubation period. These were the only crystals obtained, despite extensive attempts to crystallize this protein under these conditions or in alternative wide-ranging screening conditions. The crystals obtained were prepared for x-ray diffraction analyses by transfer to well solution, supplemented with 30% glycerol, and then flash-frozen in liquid nitrogen. Crystallographic data were collected on beamline 7-1 on an ADSC Q315R CCD detector, with an x-ray wavelength of 1.0971 Å, at the Stanford Synchrotron Radiation Lightsource. Data frames were collected using 0.5° oscillations over a 180° range at 100 Kelvin. Observed spot intensity profiles had minimal backgrounds and were well defined. Data processing was carried out with MOSFLM (44) and scaled using Scala from the CCP4 Programme Suite (45). The best crystal diffracted to 3.58 Å resolution. A molecular replacement solution was achieved using an electron microscopy model of eye lens αB-crystallin (PDB ID: 2YGD; RCSB Protein Data Bank, rcsb.org) (18), modified by CHAINSAW (45) and using the resultant polyalanine starting model in PHASER (46). Refinement was achieved by repeated alternating cycles of automated refinement in REFMAC5.8 (45) and manual building in COOT (47). REFMAC parameters used in refinement included local noncrystallographic symmetry restraints, map sharpening, and jelly-body refinement. We included an intensity-mode twin refinement in REFMAC5.8 refinement runs given that the crystal displayed 8 twin domains, including a major hkl twin domain with a fraction of 0.317. The final model was validated using the program PROCHECK (45). Related data collection and refinement statistics are given in Supplemental Table 3, and coordinates have been deposited in the Protein Data Bank (PDB ID: 6DV5; RCSB Protein Data Bank) Molecular modeling for HSP27 oligomer Molecular Operating Environment software (MOE2010) (48) was used to complete missing loops and determine the missing side chain orientations in the x-ray structure of HSP27 oligomer. MOE was also used to identify the serine phosphorylation sites on the HSP27 24-meric model. The binding mode of IVM within the phosphorylation site was identified by docking with GLIDE (49). All molecular dynamic (MD) simulations were carried out using the GPU implementation of pmemd (50) in the AMBER15 (51) package and were executed on Helios GPU cluster administered by Compute Canada. Detailed MD simulation protocols are provided below. MD simulation of HSP27 24-mer and IVM-bound HSP27 24-mer complex In order to validate the structural integrity of the predicted IVM interaction, we conducted all-atom explicit solvent MD simulation of HSP27 24-mer with 12 IVM. Structure preparation for molecular docking and MD simulation. The full HSP27 24-mer structure (Figure 1E) was constructed from the resolved x-ray structure with a homology modeling approach using the MOE suite of programs (52), and IVM was docked to the identified binding site between the NTDs using the GLIDE module in the Schrödinger suite of programs (53, 54). MD simulation protocol. All MD simulations were carried out using the GPU implementation of pmemd (50) in AMBER15 (51). ff14SB (55) was used for the HSP27 oligomer, and GAFF (56) with partial charges generated by RESP (57) was used for IVM. The solute (HSP27 oligomer or HSP27 oligomer–IVM complex) was solvated with TIP3P water molecules (58, 59) within the cubic box, ensuring a solvent shell of at least 15 Å around the solute. The system was neutralized with sodium ions. As described in the main text, the docking-predicted binding pose of IVM interacts with the NTDs of HSP27 dimers in the phosphorylation pocket (Figure 1H), where 2 of the 3 phosphorylatable serines, S78 and S82, are located. Three independent MD simulation runs were performed for 300 nanoseconds each at ambient conditions and showed that the IVM stayed well in the predicted phosphorylation pocket on the HSP27 24-mer. The average root mean square deviation (RMSD) of protein backbone over 3 runs (bold lines) and the deviation of each RMSD from their average (vertical bars) were calculated as shown in Supplemental Figure 1F. Fluorescence resonance energy transfer Fluorescence resonance energy transfer (FRET) assays were performed in Corning 384-well black flat-bottom plates. Twenty microliters of buffer (10 mM Tris, pH 8.4), 5 μL inhibitor (200 μM), 10 μL Rhodamine Red–HSP27 (Thermo Fisher Scientific) (45 nM), 10 μL QSY21-HSP27 (0.675 μL) was successively added to wells. Samples were incubated for 20 minutes at 37°C before fluorescence measurements using TECAN INFINITE M200PRO, with an excitation wavelength of 515 nm and an emission wavelength of 590 nm. Fluorescence polarization After FRET measurements, 10 μL of WT-HSP27 (2.475 μM), 10 μL of insulin-FITC (1.56 μM), and 10 μM TCEP (75 mM) were added to appropriate wells with a 2-minute equilibration period between each addition. The plate was then incubated at 42°C for 20 minutes, and fluorescence polarization (FP) measurements were taken (TECAN INFINITE F500, excitation wavelength 485 nM and emission wavelength 535 nM). Aggregation assay HSF1–/– MEF cells were lysed using 25 mM Tris (pH 7.4), 150 mM NaCl, 1 mM EDTA, 1% NP-40 lysis buffer. BSA 100 g/L and 1 M DTT were prepared in advance and thawed at room temperature. In each sample well of a Corning 96-well plate, 20 μL of BSA, 100 μL of cell lysates at 1 mg/mL, and 1 μL of DMSO or IVM with or without 1 μL of 1 mg/mL WT-HSP27 (HSP70 or HSP90) were successively added, and PBS was used to complete the total volume of 320 μL. After 5 minutes of incubation at room temperature, the plate was subjected to a 3-hour kinetic cycle with reading every 5 minutes at 340 nM at 42°C. Oligomerization assay A 12-step glycerol gradient ranging from 3.33% to 40% was constructed with 3.33% stepwise increments with total volume of 9 mL. Cell lysates were prepared in 25 mM HEPES (pH 7.4), 1 mM EDTA, 3.33% glycerol, 0.1 mM PMSF, 1 mM DTT and sonicated for 10 seconds. An equal amount of proteins, measured by bicinchoninic acid assay analysis, were loaded on a glycerol gradient and subjected to ultracentrifugation for 18 hours at 36,000 rpm (160,000 g) using a SW41Ti swing bucket rotor. Three-hundred-microliter fractions were collected into 96-well plates using a minipump and stored at –30°C for Western blot analysis. Trypsin degradation assay HSP27 and DTT were dispersed into 37°C PBS at a final concentration of 1 μM HSP27 and 20 mM DTT and allowed to equilibrate in the presence or absence of 5× (wt/wt) excess IVM for 5 minutes before addition of trypsin to a final concentration of 0.126 μM. Trypsinolysis reaction was stopped at different time points by taking 100-μL aliquots and mixing it with 2 μL of 10 M hydrochloric acid. Samples were then flash-frozen and stored at –80°C until Western blotting analysis. Actin pyrene assay HSP27 was incubated at 42°C with or without IVM. Unlabeled actin was then added to HSP27 with or without IVM and incubated for 3 minutes at 42°C. HSP27 solutions with or without IVM were added separately to pyrene-labeled actin (10:1 unlabeled to labeled actin ratio). Subsequently, the exchange buffer (10 mM EGTA, pH 8, 1 mM MgCl2) was added, and the samples were balanced using buffer A (2 mM Tris-HCl, pH 8, 200 μM ATP, 0.5 mM DTT, 0.1 mM MgCl2) and transferred into a cuvette with a final actin concentration of 4 μM. Fifty microliters of 10× KMEI buffer (500 mM KCl, 100 mM imidazole, pH 7.0, 10 mM MgCl2, 10 mM EGTA) was added to each sample. The pyrene intensity was measured by fluorometry at an excitation wavelength of 365 nm and an emission wavelength of 407 nm. Actin alone was used as control. Immunofluorescence Prostate and lung cancer cells were plated on coverslips in 12-well plates, treated with IVM for 24 hours, and exposed to heat shock (42°C for 1 hour, no recovery time). Cells were then fixed in 4% paraformaldehyde, permeabilized with 0.5% Triton, washed, blocked in PBS (3% BSA), and incubated with the anti-HSP27 primary antibody. Antigens were visualized using Alexa Fluor 488–conjugated antibodies. Nuclei were stained with DAPI VECTASHIELD mounting reagent (Vector Laboratories). Images were acquired on a Zeiss LSM-780 confocal microscope at ×63 magnification. Cancer cell lines Lung (A549 [EGFR wild-type and KRAS mutated: G12S], HCC-827 [EGFR mutated: delE746-A750]), breast (BT-474 and MCF7), prostate (22RV1, PC3), and colorectal (HCT-116, SW48) cancer cells were purchased from the American Type Culture Collection. T24 bladder cancer cells were provided by the Pathology Core of the Bladder Cancer Specialized Program of Research Excellence at MD Anderson Cancer Center (Houston, Texas, USA). Parental and HSF1–/– MEF cells were a gift from I.J. Benjamin, University of Utah (Salt Lake City, Utah, USA). Resistant HCC-827 cell lines were developed by maintenance of HCC-827 parental cells in 500 nM erlotinib for 3 months. Enzalutamide-resistant M49F cells were selected from LNCaP cells in our laboratory. Cells were maintained in RPMI 1640 medium (Invitrogen Life Technologies) containing 10% FBS (Invitrogen Life Technologies) in the presence of enzalutamide (10 μM) (11). Docetaxel-resistant PC3 cells were established in our institution, as previously reported (60). LNCaP cells were provided by Leland W.K. Chung (MD Anderson Cancer Center). PC3, MEF, MCF7, and BT-474 cell lines were maintained in DMEM (Invitrogen). LNCaP, 22RV1, A549, and HCC-827 cells were maintained in RPMI 1640 (Invitrogen Life Technologies) supplemented with 10% FBS. SW48 cells were grown in Leibovitz’s L-15 medium supplemented with 10% FBS. T24 and HCT-116 cell lines were maintained in McCoy’s 5a Medium with 5% FBS. All lines were tested for mycoplasma contamination regularly. Genomic sequencing was done to confirm cell line identity. Reagent and antibodies Erlotinib was purchased from LC Laboratories, docetaxel from Sanofi-Aventis, enzalutamide from Shanghai Haoyuan Chemexpress, and IVM from Aikon. Cetuximab and trastuzumab were obtained from the BC Cancer Agency, Vancouver Centre (Vancouver, British Columbia, Canada); polymeric micellar paclitaxel was supplied by Pharmaceutical Sciences; and 17AAG was purchased from AG Scientific Inc. IVM B1a and emamectin were obtained from AbovChem, milbemycin from Cayman Laboratory, and eprinomectin, abamectin, selamectin, and doramectin from AK Scientific. The primary antibodies used in the study were anti-HSP27 (SPA-803), anti–p-HSP27 (S78) (SPA-523), and anti–p-HSP27 (S15) (SPA-524-F) from Enzo; anti–p-HSP27 (S82) (2404S), anti-EGFR (4267S), anti–p-EGFR (Tyr1068) (3056S), anti–p-ERK (Thr202/Tyr204) (9107S), anti-AKT (9102S), anti–p-AKT (S473) (4060S), anti-PARP (9541S), anti–phospho–p38 MAPK (Thr180/Tyr182) (4511S), and anti–cleaved caspase-3 (9661) from Cell Signaling Technology; anti-SHPTP1 (ab 32559) from Abcam; anti-vinculin (V9131) and anti–β-actin (A3853) from Sigma-Aldrich; and anti-AR (441), anti–lamin B1 (sc-377001), anti-GAPDH (sc-137179), anti-CREB2 (ATF4) (sc-390063), and anti-PERK (sc-377400) from Santa Cruz Biotechnology. E-cadherin (610181) was from BD Biosciences. AR-V7 (31-1109-00) was from RevMAb Biosciences. Secondary antibodies were anti-mouse IgG–HRP (sc-2314) and anti-rabbit IgG–HRP (sc-2313) from Santa Cruz Biotechnology; and anti-mouse 800 (926-32212), anti-mouse 680 (926-68072), anti-rabbit 800 (926-32213), and anti-rabbit 680 (926-68073) from LI-COR. Secondary antibodies for immunofluorescence staining were donkey anti-mouse–Alexa Fluor 594 (A-21203) and donkey anti-rabbit–Alexa Fluor 488 (A-21206) purchased from Invitrogen Life Technologies. Transfection Cells were transfected with siRNA and apatorsen as described previously (61). The sequence of HSP27 siRNA corresponds to the human HSP27 site (5′-GUCUCAUCGGAUUUUGCAGC-3′; Dharmacon). A scrambled siRNA (5′-CAGCGCUGACAACAGUUUCAU-3′; Dharmacon) was used as a control for RNAi experiments. HSP27 ASO and scrambled (ScrB) control oligonucleotide sequences were manufactured by Ionis Pharmaceuticals and supplied by OncoGenex Technologies. The sequence of apatorsen corresponds to the human HSP27 translation initiation site (5′-GGGACGCGGCGCTCGGTCAT-3′). pHR-CMV empty and wild-type HSP27 vectors (WT) were generated and used as previously described (61). Immunoprecipitation and immunoblot analysis Whole-cell extracts were obtained by lysing of the cells in an appropriate volume of ice-cold RIPA buffer composed of 50 mmol/L Tris-HCl (pH 7.4), 150 mmol/L NaCl, 0.5% sodium deoxycholate, 1% NP-40, 0.1% SDS containing 1 mmol/L Na3VO4, 1 mmol/L NaF, 1 mmol/L PMSF, and protease inhibitor cocktail tablets (Complete, Roche Applied Science). Cellular extracts were clarified by centrifugation at 13,000 g for 10 minutes, and protein concentration was determined by a BCA protein assay kit (Thermo Fisher Scientific). Thirty micrograms of the proteins’ extracts were boiled for 5 minutes in SDS sample buffer, separated by SDS-PAGE, and transferred onto a nitrocellulose membrane. Membranes were probed with dilutions of primary antibodies listed above, followed by incubation with either HRP-conjugated or fluorescent secondary antibodies. After washing, proteins were visualized by either a chemiluminescent detection system (GE Healthcare) or an Odyssey Imaging System (LI-COR). For coimmunoprecipitation the soluble proteins were precleared with protein A/G-Sepharose beads for 1 hour. Precleared lysates were incubated overnight with 5 μg antibody or IgG at 4°C with constant rotation. A/G beads were added for 1 hour, and after centrifugation, the pellet was washed twice. Proteins were eluted from the beads by incubation of the immunoprecipitates with 60 μL of 2× SDS sample buffer (24 mM Tris, pH 6.8, 10% [vol/vol] glycerol, 0.8% [vol/vol] SDS, 6 mM 2-mercaptoethanol, and 0.04% bromphenol blue) at 90°C for 3 minutes. Nuclear/cytoplasmic fractions were extracted using the CelLytic NuCLEAR Extraction Kit (Sigma-Aldrich) according to the manufacturer’s protocol. Molecular weight markers are specified in all the blots. Migration (wound healing) assay HCC-827 and LNCaP cells were plated in coated 96-well ImageLock plates (Essen Bioscience) at 2 × 104 cells per well. After 24 hours the wound was made in each well using Woundmaker (Essen Bioscience), and the cells were treated in the presence or absence of IVM. The wound closure was monitored for an additional 42 hours with an IncuCyte machine every 6 hours. Relative wound density was determined using IncuCyte software. Biolayer interferometry assay Direct reversible interactions between IVM and purified HSP27 were quantified by biolayer interferometry (BLI) using OctetRED (ForteBio). HSP27 in fusion with an N-terminal (His) 6-avidin (GLNDIFEAQKIEWHE) tag was expressed in E. coli BL21-DE3 cells cotransformed with pBirACm plasmid, which expresses a biotin ligase leading to the biotinylation of the avidin tag on HSP27. Cells were grown in LB medium at 37°C and then induced with 0.5 mM isopropyl-β-d-thiogalactoside in the presence of 0.15 mM biotin. Purified protein at 200 μg/mL was bound to the streptavidin sensors overnight at 4°C in gel filtration buffer (20 mM Tris-base pH 8.4, 100 mM NaCl, 0.2 mM TCEP, 0.1 mM PMSF). The assay was performed at 25°C in the dialysis buffer supplemented with 1% DMSO, which was also used as our reference measurement. Enzalutamide was used as negative control. In vitro kinase assay For in vitro kinase reactions, 35 μM wild-type HSP27 was incubated with either DMSO or 350 μM IVM for 1 hour in reaction buffer (20 mM Tris, pH 8, 150 mM NaCl, 5% glycerol, 0.2 mM TCEP, 17 mM MgCl2, 350 μM ATP, 13% DMSO). Subsequently, 35 ng/mL of MAPKAP2 kinase was added to each mixture and incubated at 30°C. Samples were taken at t = 0 and 24 hours, and the kinase reaction was stopped by addition of 5× protein loading dye and boiling for 5 minutes. Phosphorylation was determined using the HSP27 S78 antibody by Western blotting. Sample extraction and liquid chromatography–mass spectrometry analysis Serum samples from the in vivo studies were thawed and 20 μL transferred to individual Eppendorf tubes, followed by addition of internal standard (IS; 4 μL of 2 μg/mL abamectin) and 44 μL of PPT (methanol/acetonitrile, 25:75). Samples were vortexed, precipitated protein sedimented (5 minutes, 20,000 g), and clarified supernatant transferred to liquid chromatography vials. Standards were prepared in a similar fashion with IVM/IS–spiked blank mouse serum; parallel standards made up in solvent only were used to characterize any matrix effects. Tissue samples (200–400 mg) were weighed and 2 volumes water added, followed by homogenization using a Precellys tissue homogenizer (Bertin Technologies) with three 45-second cycles at 6000 rpm. Extraction was similar to that of serum but using 50 μL homogenate, 10 μL IS, and 110 μL PPT, and spiked serum samples were used for calibration. An Acquity UPLC System coupled with an eLambda Photodiode Array (PDA) and a Quattro Premier mass spectrometer (Waters) was used for analysis with a 100-mm BEH C18, 1.7-μm column (Waters) and an acetonitrile gradient (0–0.5 minutes, 88%; 0.5–1.5 minutes, 88%–98%; 1.5–3.5 minutes, 98%; 3.5–3.6 minutes, 98%–88%; 0.3 mL/min) used for separation (5.5-minute run length; 0.1% formic acid present throughout). Mass spectrometry data were collected in electrospray positive slightly below unit resolution with the following instrument parameters: capillary, 3 kV; extractor and RF lens, 3 V and 0.2 V; source and desolvation temperatures, 120°C and 350°C; desolvation and cone (N2) flow, 950 L/h and 50 L/h; collision gas (Ar) flow, 0.15 mL/min (8–3-meter bar). Compounds were detected by multiple reaction monitoring with m/z 892.7>307.3 and 892.7>569.3 for IVM and m/z 890.7>305.3 and 890.7>567.3 for Adamectin (20/24 V, 20/15 V, and 20/25 V, 20/12 V, cone/collision volt combinations, respectively) with 0.1 seconds dwell each. Retention times for IVM and Adamectin were 2.6 minutes and 1.7 minutes, respectively. Extracted chromatograms of 254 nm were used for both IVM and Adamectin using the PDA data (210–500 nm). Data were processed with Quanlynx (Waters) using Adamectin-normalized calibration for mass spectrometry and external calibration for PDA and exported to Excel for further analysis. Calibration standards ranged from 0.4 to 1200 ng/mL (6 points, extract levels). A linear fit was used, resulting in r2 > 0.98, and deviation from nominal < 15% above 1.6 ng/mL. OD data correlated well at higher concentrations. Ion suppression in both serum and tumor extracts was on the order of 30%. Quantitative reverse transcriptase PCR RNA extraction and reverse transcriptase PCR (RT-PCR) were carried out as described previously (42). Real-time monitoring of PCR amplification of cDNA was carried out using the following primer pairs and probes: EGFR Hs01076090_m1, CDH1 Hs1023894_m1, and GAPDH Hs03929097_g1 (Applied Biosystems), on the ABI PRISM 7900 HT Sequence Detection System (Applied Biosystems) with a TaqMan Gene Expression Master Mix (Applied Biosystems). Target gene expression was normalized to GAPDH levels in the respective samples as an internal control. The results are representative of 3 independent experiments. Doxycycline-inducible AR-V7 PC3_3TKNLuc system Doxycycline-inducible AR-V7 construct and probasin-based ARR3tk-Nanoluciferase reporter construct (PC3V7_3TKNLuc) were generated as previously reported (62). Nanoluciferase reporter assays were performed by culturing of cells in DMEM plus 5% charcoal-stripped serum medium in black 96-well plates. AR-V7 expression was induced by the addition of 200 ng/mL doxycycline with different doses of IVM for 24 hours, and cells were lysed with Nanoglo (Promega) lysis buffer/substrate for 5 minutes at room temperature. Luminescence was measured by a TECAN m200pro luminometer. In vitro cell growth assay Cells were plated in 96-well plates and treated with IVM alone or in combination with erlotinib, cetuximab, trastuzumab, or docetaxel. After 3 days, cell viability was assessed using crystal violet assay. Absorbance was determined with a microculture plate reader (Becton Dickinson Labware) at 590 nm. Changes in cell viability were calculated as percentage relative to controls. Each assay was performed in triplicate. Tissue microarray construction and immunohistochemistry We created the tissue microarray by taking multiple 1-mm cores from matching marked non-necrotic areas of paraffin blocks of xenograft tumors using a semiautomated tissue microarrayer (TMArrayer, Pathology Devices) attached to a Leica M50 stereo microscope. Staining was conducted using an automated staining platform (Ventana Discovery XT; Ventana Medical Systems) with an enzyme-labeled biotin streptavidin system and solvent-resistant DAB Map Detection Kit. TUNEL staining was conducted by Ventana Autostainer using protease for antigen retrieval. The mixture of terminal transferase, digoxigenin-11-dUTP, and dATP was added, followed by biotinylated anti-digoxigenin IgG, and the DAB Map Detection Kit was used. All stained slides were digitalized with the SL801 autoloader and Leica SCN400 scanning system (Leica Microsystems) and were subsequently stored in the SlidePath digital imaging hub (DIH; Leica Microsystems) of the Vancouver Prostate Centre. Using the Aperio Image Analysis IHC (Leica Biosystems), our pathologist selected the area of interest, defined the parameter, optimized the level of intensity, and selected Positive Pixel Count algorithm for each biomarker. HSP27 antibody (SPA-803, Stressgen) was used at a concentration of 1:3000, p-HSP27 antibody (9308, Cell Signaling Technology) at 1:50, and their use for immunohistochemistry was validated as previously reported (11). The EGFR and p-EGFR antibodies (4267 and 3777, Cell Signaling Technology) were used at a concentration of 1:50 and were validated by Western blotting in NSCLC and MEF cell lines, as shown in Figure 3, A–C and F. SHPTP1 antibody (32559, Abcam) was validated in liver and lymphoid tissue, as negative and positive controls, respectively, and was used at a concentration of 1:500. In situ proximity ligation assay Proximity ligation assay was performed to detect the interaction between HSP27 and p-EGFR and HSP27 and EGFR. To visualize the bound antibody pairs, the Duolink Detection Kit (Duo92008) with proximity ligand assay (PLA) PLUS and MINUS probes for mouse and rabbit (Olink Bioscience) was used, according to the manufacturer’s description. Cell slides were mounted with Duolink Brightfield Mounting Medium (Olink Bioscience). SHPTP1 phosphatase assays In vitro SHPTP1 tyrosine phosphatase assays were performed using the Malachite Green Phosphatase Assay (Upstate Biotechnology) with phosphopeptide (RRLIEDAEpYAARG) as a substrate from Promega. Briefly, cells were lysed in NP-40 lysis buffer and incubated with anti-SHPTP1 overnight at 4°C followed by 1 hour of incubation with protein A/G-Sepharose beads. The immune complexes were washed 3 times with lysis buffer without phosphatase inhibitor and resuspended in phosphatase buffer containing the phosphopeptide. The reaction mixtures were incubated for 30 minutes at room temperature followed by the addition of Malachite Green solution. Absorbance was measured in a spectrophotometer at 620 nm. Phosphate release was determined by comparison of absorbance to the phosphate standard. Animal manipulation and assessment of in vivo tumor growth For the HCC-827 in vivo experiment, 6- to 8-week-old female athymic mice (Harlan Sprague Dawley Inc.) were inoculated with 2 × 106 tumor cells s.c. in the left flank via a 27-gauge needle under isoflurane anesthesia. When tumors reached 150–200 mm3, mice were randomized to receive 15 mg/kg erlotinib, 10 mg/kg IVM, or the combination of both drugs (formulated in 0.5% methylcellulose, 0.1% Tween-80) administered orally 3 days a week. For the LNCaP in vivo experiment, 2 × 106 LNCaP cells were inoculated s.c. into the left and right flanks of 6- to 8-week-old male athymic nude mice. The animals were surgically castrated when the PSA reached 50 ng/mL or the tumor volume 300 mm3 and were subsequently randomized to receive vehicle or IVM treatment, using the same schedule protocol described for the HCC-827 study. For the 22RV1 in vivo experiment, 2 × 106 22RV1 cells were inoculated s.c. into the left flank of 6- to 8-week-old male athymic nude mice. When the tumor volume reached 300 mm3, the animals were randomized to receive vehicle or IVM, as reported above. Six- to eight-week-old male athymic nude mice were inoculated with 6 × 106 docetaxel-resistant PC3 cells s.c. in the left flank. When tumors reached 150 mm3, mice were randomly selected for treatment with micellar paclitaxel (0.5 mg, 3 times per week) administered i.v. every other week, 10 mg/kg IVM by gavage 3 times a week, or the combination of both. For all the in vivo experiments, tumor volume measurements were performed twice weekly and calculated by the formula length × width × depth × 0.5236. Data points were expressed as average tumor volume ± SEM. Statistical analysis For the in vitro experiments, unpaired 2-tailed Student’s t test and ANOVA followed by Bonferroni’s multiple-comparisons test were used. For the in vivo analysis, 1-way ANOVA test with Tukey’s correction was used to compare 3 or more groups and unpaired t test with Welch’s correction to compare 2 groups. GraphPad Prism 6 software was used to calculate the statistical significance. The threshold of statistical significance was set at *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001. Study approval All animal procedures were performed according to the guidelines of the Canadian Council on Animal Care and with appropriate institutional certification. Animal studies were approved by the Animal Care Committee of the University of British Columbia (protocol number A18-0118).