NKX3.1 localization to mitochondria suppresses prostate cancer initiation

Abstract

Mitochondria provide the front-line of defense against the tumor-promoting effects of oxidative stress. Here we show that the prostate-specific homeoprotein, NKX3.1, suppresses prostate cancer initiation by protecting mitochondria from oxidative stress. Integrating analyses of genetically-engineered mouse models, human prostate cancer cells, and human prostate cancer organotypic cultures, we find that, in response to oxidative stress, NKX3.1 is imported to mitochondria via the chaperone protein, HSPA9, where it regulates transcription of mitochondrial-encoded electron transport chain (ETC) genes, thereby restoring oxidative phosphorylation and preventing cancer initiation. Germline polymorphisms of NKX3.1 associated with increased cancer risk fail to protect from oxidative stress or suppress tumorigenicity. Low expression levels of NKX3.1 combined with low expression of mitochondrial ETC genes are associated with adverse clinical outcome, whereas high levels of mitochondrial NKX3.1 protein are associated with favorable outcome. This work reveals an extranuclear role for NKX3.1 in suppression of prostate cancer by protecting mitochondrial function.

Statement of Significance

Our findings uncover a non-nuclear function for NKX3.1 that is a key mechanism for suppression of prostate cancer. Analyses of the expression levels and sub-cellular localization of NKX3.1 in patients at risk of cancer progression may improve risk assessment in a precision prevention paradigm, particularly for men undergoing active surveillance.

Introduction

Prostate cancer is the most prevalent type of cancer and a leading cause of cancer-related death in men (1,2). Nowadays most prostate cancers are diagnosed with early stage disease, which is usually not life-threatening (5-year survival of >90%)(3,4). Consequently, it has become common practice to monitor men by “active surveillance”, which avoids treatment unless there is evidence of progression. While active surveillance minimizes overtreatment, it may miss the opportunity for early intervention of tumors that are likely to progress to advanced prostate cancer, which, unlike locally-invasive disease, is often lethal (5-year survival of <30%) (3,4).

A distinguishing feature of prostate cancer is its long latency, since pre-invasive lesions, termed prostatic intraepithelial neoplasia (PIN), are common in young adults whereas clinically-detectable disease is usually not manifested until the sixth decade or later (2,5,6). Risk factors for prostate cancer include aging and inflammation, while incidence and aggressiveness are influenced by genetic factors and race (2,6,7). Given its long latency, men diagnosed with or at risk for prostate cancer may benefit from early intervention to delay progression, which underscores the importance of elucidating mechanisms of cancer initiation that may provide opportunities for precision prevention (8,9).

A key factor associated with prostate carcinogenesis is oxidative stress (10-12), which leads to excessive production and accumulation of reactive oxygen species (ROS) (13-15). Most cellular ROS is produced by mitochondria as a byproduct of cellular respiration (15-17). In normal cellular contexts, ROS levels and oxidative phosphorylation (OXPHOS) are tightly balanced, which is important for mitochondrial homeostasis. However, aberrant oxidative stress may lead to excess ROS and thereby impaired mitochondrial function (13,14). The electron transport chain (ETC), which generates mitochondrial ROS, is located on the inner mitochondrial membrane and is comprised of several protein complexes whose subunits are encoded by both nuclear and mitochondrial genes (18). Consequently, communication between the nucleus and mitochondria is essential for sustaining ETC function and appropriate ROS levels in response to oxidative stress (19).

Although the role of ROS in cancer is complex (13), substantial evidence indicates that the prostate is particularly susceptible to elevated ROS as a consequence of oxidative stress associated with inflammation, aging, or other factors (11,12,20). Furthermore, alterations of mitochondrial function or mutational burden, which may impact ROS levels or response to oxidative stress, are prevalent in prostate cancer and associated with adverse disease outcome (21-25). Among genes that have been attributed to protect against oxidative stress in prostate, is NKX3.1, a prostate-specific homeobox gene that regulates prostate differentiation and suppresses prostate cancer initiation (26). In mice, loss of function of Nkx3.1 leads to PIN, which, in collaboration with loss of other tumor suppressor genes, progresses to adenocarcinoma (27-30). In humans, allelic imbalance of NKX3.1 is among the most prevalent and earliest genetic alterations in prostate cancer, while reduced expression of NKX3.1 is associated with increased risk of disease aggressiveness, particularly among African-American men (31-37).

NKX3.1 is expressed in the prostatic epithelium, where it functions to safeguard prostatic specification and maintenance of luminal prostatic stem cells, and to protect against DNA damage and inflammation (38-46). Since NKX3.1 encodes a homeoprotein, which are known regulators of nuclear gene transcription, its functions in prostate have long been attributed to regulation of nuclear gene expression. Here, we uncover a non-nuclear role for NKX3.1 in mitochondria, where it regulates expression of mitochondrial genes to promote mitochondrial homeostasis and suppress tumorigenesis. Notably, we find that NKX3.1 expression levels and sub-cellular localization are prognostic of clinical outcome in prostate cancer patients. We propose that analyses of NKX3.1 expression and localization may enhance precision monitoring of prostate cancer patients to identify those at risk of progression.

Results

Oxidative stress accelerates prostate cancer initiation and impairs mitochondria in Nkx3.1 mutant mice

Based on our previous study showing that loss-of-function of Nkx3.1 in mouse prostate leads to increased expression of genes associated with oxidative stress (40), we performed RNA sequencing comparing prostate tissues from Nkx3.1 wild-type (Nkx3.1^+/+) or mutant (Nkx3.1^−/−) mice (27) with human RWPE1 prostate cells expressing or lacking exogenous NKX3.1 (39) in normal contexts or in response to oxidative stress (Datasets 1, 2). We found that NKX3.1-dependent biological pathways are highly conserved in mouse and human prostate, particularly following oxidative stress (Supplementary Fig. S1A,B; Datasets 3,4). Conserved pathways include those associated with regulation of oxidative stress, mitochondrial function, and glucose metabolism (Supplementary Fig. S1C,D). Therefore, we have investigated the functions of NKX3.1 in response to oxidative stress in both mouse models and human prostate cells.

First, we studied the consequences of oxidative stress for prostate carcinogenesis in Nkx3.1 mutant mice, which develop PIN indicative of pre-invasive prostate cancer by 12 months of age (Fig. 1A; Supplementary Table S1) (27,28). We treated adult Nkx3.1^+/+ or Nkx3.1^−/− mice continually for 1-9 months with paraquat (N,N′-dimethyl-4,4′-bipyridinium dichloride [(C₆H₇N)₂Cl₂), a redox-active compound that produces superoxide anions (47) (Fig. 1A-H; Supplementary Fig. S2A-E). As a control, we analyzed seminal vesicle (Supplementary Fig. S2F-J, Supplementary Table S1), which, like prostate, is an androgen-regulated male secondary sexual organ, but unlike prostate does not express Nkx3.1 and rarely develops cancer (27,39).

Figure 1. NKX3.1 protects against mitochondrial oxidative stress.

(A-H) Analyses of Nkx3.1 mutant mice. (A) Strategy. Nkx3.1 mutant (Nkx3.1^−/−), but not wild-type (Nkx3.1^+/+) mice develop PIN by 12 months of age. Cohorts of mice received paraquat (10mg/kg/day in drinking water) or vehicle (water alone) starting at 3 months of age and were sacrificed at 4 months to measure ROS levels or 12 months for phenotypic analysis. (B) Quantification of DHE fluorescence from the anterior prostate of Nkx3.1^+/+ or Nkx3.1^−/− mice treated with paraquat (Par) or vehicle (Veh) for 1 month (n=9-14 mice/group). (C) Histopathology showing hematoxylin and eosin (H&E) staining and immunostaining for Nkx3.1 and γH2AX of anterior prostate from mice treated with paraquat or vehicle for 9 months and analyzed at 12 months of age. Scale bars represent 50μm (low power) or 20μm (high power). Shown are representative images from analyses of 18-28 mice per group. (D) Quantification of the PIN phenotype showing the relative percentage of low-grade PIN (PIN1/2) and high-grade PIN (PIN3/4) in anterior prostate. Data show the summary from analysis of 18-28 mice/group and are expressed as the mean percentage ± SD of the control; P values were calculated using chi-squared (χ²) test. ns, not significant. (E-G) Electron microscopy of mitochondria from anterior prostate of Nkx3.1^+/+ and Nkx3.1^−/− mice treated with paraquat (10 mg/kg/day) or vehicle for 9 months and analyzed at 12 months of age. (E) Representative micrographs; scale bars represent 100nm. (F) Quantification of relative mitochondrial area. (G) Quantification of relative number of mitochondria. Panels F and G represent analyses of 60+ images/mouse and 4 mice/group and expressed as the mean percentage ± SD. (H) Quantification of mitochondrial membrane potential as detected by TMRE fluorescence from mice treated with paraquat (10 mg/kg/day) or vehicle for 9 months and analyzed at 12 months of age. (I-W) Analyses of human prostate cells in culture. (I) Strategy. LNCaP, BPH1, or RWPE1 were treated with paraquat or MitoParaquat to induce general cellular or mitochondrial-specific reactive oxygen species (ROS), respectively, alone or together the mitochondrial-specific anti-oxidant, MitoQ. Production of general cellular ROS or mitochondrial ROS was measured using the dyes indicated. (J-W) Oxidative stress was induced by treating cells with paraquat (100μM for 24 hours), hypoxia (1% pO₂ for 24 hours) or hydrogen peroxide (200μM for 6 hours). (J, K) Western blot analyses of total protein lysates in LNCaP cells expressing 2 independent shRNA for NKX3.1 (shNKX3.1#1, shNKX3.1#2) or the control (shControl) (J) and RWPE1 cells expressing NKX3.1 or the control vector (K). Note that subsequent data show shNKX3.1#1. (L, M) Western blot analyses of total protein lysates in LNCaP cells expressing shRNA for NKX3.1 (shNKX3.1) or the control (shControl) (L) and NKX3.1-expressing (or control) RWPE1 cells (M) treated with paraquat (100μM for 24 hours). γH2AX is a marker of DNA damage. (N-R) Analysis of LNCaP cells. (N) Representative histograms from ROS analyses. (O and P) Quantification of mean ROS intensity as detected by DHE (O) or MitoSOX (P) production. (Q) Quantification of mean mitochondrial mass/density as detected by MitoTracker. (R) Quantification of mitochondrial membrane potential as detected by TMRE fluorescence. (S) Representative histograms from ROS analyses. (T, U) Quantification of mean ROS intensity as detected by DHE (T) or MitoPY1 (U) production. (V) Quantification of mean mitochondrial mass/density as detected by MitoView. (W) Quantification of mitochondrial membrane potential as detected by TMRE fluorescence. Panels N-W show representative data from 3 independent experiments; each experiment was done in triplicate or quadruplicate (9-12 independent samples/group). Unless otherwise indicated, shown is the mean ± SD; P values were calculated using two-sample unpaired Welch t-test. ns, not significant. See also Supplementary Figures S1-S5 and Supplementary Tables S1- S4.

After 1 month (i.e., in 4 month-old mice), paraquat-treated Nkx3.1^−/− prostates displayed increased ROS, which was not observed in Nkx3.1^+/+ prostates, or seminal vesicles of either genotype (Fig. 1B, Supplementary Fig. S2A,B,G). Further, after 9 months (i.e., in 12 month-old mice) we observed a significant acceleration of PIN in all prostatic lobes of Nkx3.1^−/− mice, but not in prostates of Nkx3.1^+/+ mice or seminal vesicles from either genotype (Fig. 1C,D; Supplementary Fig. S2C,F). Acceleration of prostate carcinogenesis was evident by the significant increase in high-grade PIN (PIN3/4) compared with low-grade PIN (PIN1/2) in paraquat-treated versus control Nkx3.1^−/− prostate (Fig. 1D; Supplementary Table S1). This was accompanied by increased DNA damage, as evident by immunostaining for γH2AX (Fig. 1C). These phenotypes were not due to general paraquat toxicity, since no other major organs were grossly affected by long-term paraquat treatment (Supplementary Fig. S3), consistent with previous studies showing that prostate is particularly susceptible to stressful stimuli (46).

To ask whether oxidative-stress resulted in abnormalities of mitochondrial morphology we performed electron microscopy (EM). These analyses revealed a significant reduction in number and area of mitochondria in the paraquat-treated Nkx3.1^−/− prostate, but not the Nkx3.1^+/+ prostate or in the seminal vesicle (Fig. 1E-G; Supplementary Fig. S2D,H,I; Supplementary Table S2). Aberrations of mitochondrial morphology included condensation, lysis, and membrane dissolution (Fig. 1E). We also observed a significant reduction in active mitochondria specifically in the paraquat-treated Nkx3.1^−/− prostate, evidenced by decreased mitochondrial membrane potential as measured using TMRE (tetramethylrhodamine, ethyl ester) (Fig. 1H; Supplementary Fig. S2E,J). Therefore, NKX3.1 protection against oxidative stress in vivo is associated with suppression of cancer initiation and protection of mitochondrial integrity.

NKX3.1 protects against mitochondrial ROS in cancerous and benign human prostate cells

We next investigated the functions of NKX3.1 for protection against oxidative stress in human prostate cells in vitro (Fig. 1I-W; Supplementary Figs. S4A-K, S5A-F; Supplementary Tables S3, S4). In particular, we silenced endogenous NKX3.1 in LNCaP cells (Fig. 1J), one of the few human prostate cancer cell lines that expresses endogenous NKX3.1 (41), and in BPH1 cells (Supplementary Fig. S5A), which are immortalized benign prostatic hyperplasia cells that express endogenous NKX3.1 (48). Alternatively, we expressed exogenous NKX3.1 in RWPE1 cells (Fig. 1K), an immortalized human prostate epithelial cell line with negligible levels of endogenous NKX3.1 (39). We induced oxidative stress by treating cells with paraquat and measuring accumulation of cellular superoxide (O⁻₂) or hydrogen peroxide (H₂O₂) ROS, using dihydroethidium (DHE) or 6-chloromethyl-2′,7′-dichlorodihydrofluorescein diacetate, acetyl ester (CM-H2DCFDA), respectively (Fig. 1I).

In these cancerous and benign contexts, cells expressing or lacking NKX3.1 had similar ROS levels in standard culture conditions (i.e., with vehicle) while oxidative stress was associated with increased ROS in cells lacking but not those expressing NKX3.1 (Fig. 1N-W; Supplementary Fig. S4A-K, S5A-F). This was accompanied by increased DNA damage as evident by expression of γH2AX (Fig. 1L,M). In particular, treatment of NKX3.1-silenced LNCaP or BPH1 cells with paraquat significantly increased ROS, which was not observed in the corresponding control cells (Fig. 1N,O; Supplementary Figs. S4B, S5B,C). Notably, the effects of silencing NKX3.1 in LNCaP cells were rescued by exogenous NKX3.1 (Supplementary Fig. S4C-E). Conversely, paraquat treatment resulted in increased ROS in control RWPE1 cells (which lack NKX3.1) but not in the NKX3.1-expressing cells (Fig. 1S,T; Supplementary Fig. S4H). Similar results were obtained by inducing oxidative stress with hydrogen peroxide or hypoxia (Fig. 1O,T; Supplementary Figs. S4B,H, S5C).

Given the Nkx3.1-dependent effects of oxidative stress on mitochondria in vivo, we asked whether NKX3.1 protects specifically against mitochondrial ROS (compared with general cellular ROS) by measuring mitochondrial-specific superoxide (O⁻₂) or hydrogen peroxide (H₂O₂) using MitoSOX or MitoPY1, respectively (Fig. 1I). As observed for general cellular ROS, mitochondrial-specific ROS was significantly increased following paraquat treatment in NKX3.1-silenced LNCaP and BPH1 cells, but not the corresponding control cells (which express NKX3.1) (Fig. 1P; Supplementary Figs. S5D). Conversely, oxidative stress led to increased mitochondrial ROS in control but not NKX3.1-expressing RWPE1 cells (Fig. 1U).

Induction of oxidative stress with MitoParaquat, which has a similar mode of action as paraquat but induces oxidative stress preferentially in mitochondria (49) resulted in increased ROS in cells lacking NKX3.1 (i.e., in NKX3.1-silenced BPH1 cells and control RWPE1 cells), which was fully abrogated in the corresponding NKX3.1-expressing cells (i.e., control BPH1 and NKX3.1-expressing RWPE1 cells) (Supplementary Figs. S4I, S5E,F). Furthermore, increased ROS in cells lacking NKX3.1 was rescued by the mitochondrial-specific anti-oxidant, MitoQ (50) (Fig. 1I; Supplementary Fig. S4F,J), further demonstrating that NKX3.1 protects primarily against mitochondrial ROS. In addition, oxidative stress resulted in the significant reduction of active mitochondria in cells lacking NKX3.1, as evident by measurement of mitochondrial mass and/or membrane potential (Fig. 1Q,R,V,W; Supplementary Fig. S4G,K). These findings demonstrate that NKX3.1 protects against mitochondrial ROS, which is associated with protection of mitochondrial integrity.

NKX3.1 localizes to mitochondria following oxidative stress

Although Nkx3.1 displayed the expected nuclear localization in vehicle-treated mouse prostate, its sub-cellular localization in paraquat-treated prostate did not appear exclusively nuclear (see Fig. 1C, inset). This prompted us to investigate the sub-cellular localization of NKX3.1 in response to oxidative stress. Under standard conditions (i.e., in the vehicle-treated contexts) NKX3.1 was localized primarily to nuclei in mouse prostate tissues and human prostate cells; in contrast, following oxidative stress, NKX3.1 became localized to mitochondria as well as nuclei (Fig. 2A-D; Supplementary Figs. S5G,H, S6A). This was evident by co-localization of NKX3.1 with the mitochondrial-specific marker, ATPB (ATP synthase subunit beta) as visualized by confocal imaging (Fig. 2A,C; Supplementary Fig. S5G), and western blot analysis following biochemical fractionation to isolate nuclei or mitochondria (Fig. 2B,D; Supplementary Figs. S5H, Supplementary Fig. S6A).

Figure 2. NKX3.1 localizes to mitochondria in response oxidative stress.

(A-E) Localization of NKX3.1 to mitochondria. Panels A and B, show analysis of anterior prostate from Nkx3.1^+/+ mice treated with paraquat (10mg/kg/day in drinking water) or vehicle (water alone) for 9 months. Panels C and D show analysis of LNCaP cells treated with paraquat (100μM) or vehicle for 24 hours. (A,C) Confocal images. Samples were co-stained with an antibody specific for mouse or human NKX3.1 (red) and anti-ATPB to visualize mitochondria (green); nuclei were visualized by labeling with DAPI (blue). Scale bars in Panel A represent 50μm (left and center) or 200μm (right) and in Panel C 100μm. (B,D) Western blot analyses following biochemical isolation of nuclear or mitochondrial fractions, as detected using histone H3 (H3) and ATPB, respectively. (E) Representative immuno-EM photomicrographs from LNCaP cells to detect endogenous NKX3.1 or NKX3.1-expressing RWPE1 cells to detect the exogenous protein. Cells were treated with paraquat (100μM) or vehicle (media alone) for 24 hours followed by detection of NKX3.1 (black dots) in mitochondria using an anti-NKX3.1 antibody. Scale bars represent 100nm (left and center) or 400nm (right). (F-P) Analyses of NKX3.1 polymorphisms associated with cancer risk. (F)Schematic showing the amino acid substitutions encoded by the NKX3.1 polymorphisms, Arginine 52 to Cysteine (R52C) and Threonine 164 to Alanine (T164A), or the control, Arginine 52 to Alanine (R52A). Also shown is a summary of the DNA binding, mitochondrial localization and ROS response. High level activity is indicated by (+) and no/low level activity by (−). (G-I) Analyses of RWPE1 cells expressing NKX3.1 (wildtype), T164A, R52C, or R52A treated with vehicle or paraquat (100μM) for 24 hours. (G) Western blot analyses following isolation of nuclear or mitochondrial fractions. (H and I) Quantification of mean ROS intensity, as detected by DHE (H) or MitoPY1 (I) to measure general cellular or mitochondrial ROS, respectively. (J-P) Analyses of LNCaP cells expressing shNKX3.1 (or a control shRNA) alone or with exogenous NKX3.1, T164A, or R52C. Cells were treated with vehicle or paraquat (100μM) for 24 hours. (J) Western blot analyses following isolation of nuclear or mitochondrial fractions. (K and L) Quantification of mean ROS intensity, as detected by DHE (K) or MitoSOX (L). (M) Quantification of cellular proliferation as detected by MTT absorbance. (N) Representative images of colony formation assays visualized by staining for crystal violet. (O) Quantification of colony number. (P) Quantification of Matrigel invasion assays. Unless otherwise indicated, shown are representative data from 2-3 independent experiments, each done in triplicate (9 independent samples/group) showing mean ± SD; P values were calculated using two-sample unpaired Welch t-test. ns, not significant. See also Supplementary Figures S6 and S7 and Supplementary Tables S3 and S5.

To more precisely study localization of NKX3.1 in mitochondria, we performed immuno-electron microscopy (immuno-EM) on LNCaP and RWPE1 cells (Fig. 2E). This revealed that, following oxidative stress, NKX3.1 was localized within the interior of mitochondria, where it is associated with the cristae of the inner mitochondrial membrane (Fig. 2E). Localization within the interior of mitochondria was further supported by biochemical fractionation analyses, in which NKX3.1 was resistant to proteolysis in intact mitochondria but not after disruption by sonication (Supplementary Fig. S6B).

Polymorphisms associated with increased cancer risk fail to protect against oxidative stress or suppress tumorigenicity

To gain insights regarding the relevance of these findings for NKX3.1-related cancer risk, we examined the localization and activities of two germline polymorphisms encoding NKX3.1(T164A) and NKX3.1(R52C) (Fig. 2F-L; Supplementary Fig. 6C-I; Supplementary Table S5) (51,52). NKX3.1(T164A) occurs in 14% of the population and is associated with early onset prostate cancer (52), whereas NKX3.1(R52C) is present in 11% of the population and is associated with increased cancer aggressiveness (51). NKX3.1(T164A) corresponds to a threonine-to-alanine substitution in the homeodomain that renders the protein inactive for DNA binding (39), whereas NKX3.1(R52C) corresponds to an arginine-to-cysteine substitution in the N-terminal region that does not impair DNA binding (Fig. 2F,; Supplementary Fig. S6C) (51).

NKX3.1(T164A) and NKX3.1(R52C) were located only in the nucleus of RWPE1 cells in standard culture conditions, as evident by confocal imaging and western blot analysis; however, following oxidative stress, NKX3.1(T164A) was localized to mitochondria as well as nuclei, whereas NKX3.1(R52C) was only in the nucleus (Fig. 2G, Supplementary Fig. S6D). This deficiency of mitochondrial localization of NKX3.1(R52C) was specific for the arginine-to-cysteine substitution since a protein with a substitution of arginine-to-alanine [NKX3.1(R52A)] localized to mitochondria following oxidative stress (Fig. 2G).

Interestingly, neither NKX3.1(T164A) nor NKX3.1(R52C) reduced general cellular or mitochondrial-specific ROS (Fig. 2H,I; Supplementary Fig. S6E-I), suggesting that multiple regions of NKX3.1 are required for protection against ROS. Indeed, only full length NKX3.1, but not various truncated proteins, localized to mitochondria and alleviated ROS in response to oxidative stress (Supplementary Fig. S7A-E).

To further study their localization and functions and to assess the relevance for tumorigenicity, we asked whether NKX3.1(T164A) or NKX3.1(R52C) could rescue the consequences of silencing NKX3.1 in LNCaP cells, as we had observed for native NKX3.1 (see Supplementary Fig. S4C-E). As in RWPE1 cells, exogenous NKX3.1, NKX3.1(T164A) and NKX3.1(R52C) were located in nuclei in NKX3.1-silenced in LNCaP cells in standard conditions, while following oxidative stress native NKX3.1 and NKX3.1(T164A), but not NKX3.1(R52C) localized to mitochondria (Fig. 2J). Additionally, only native NKX3.1, but neither NKX3.1(T164A) nor NKX3.1(R52C), could alleviate ROS in response to oxidative stress in NKX3.1-silenced in LNCaP cells (Fig. 2K,L). Notably, oxidative stress resulted in increased cellular proliferation, colony formation and invasion in NKX3.1-silenced LNCaP cells, which was abrogated by native NKX3.1, but not NKX3.1(T164A) or NKX3.1(R52C) (Fig. 2M-P).

Together, these findings demonstrate that protection from mitochondrial ROS as well as suppression of tumorigenicity in response to oxidative stress require NKX3.1 binding to DNA, which is impaired for NKX3.1(T164A), and localization to mitochondria, which is impaired for NKX3.1(R52C). Furthermore, these observations raise the possibility that increased risk of aggressive prostate cancer associated with these germline variants of NKX3.1 is due, at least in part, to a loss of protection of mitochondria from oxidative stress.

HSPA9 imports NKX3.1 to mitochondria in response to oxidative stress

We consistently observed that localization of NKX3.1 to mitochondria was not accompanied by its reduced nuclear expression; rather, relative levels of nuclear NKX3.1 were similar in standard conditions and following oxidative stress (see Fig. 2B,D; Supplementary Figs. S5H, S6A). Time-course analysis revealed localization of NKX3.1 to mitochondria beginning at 6 hours after oxidative stress and peaking at 24 hours, during which time the levels of nuclear NKX3.1 remained relatively constant (Supplementary Fig. S8A); this paralleled the time-course of protection from oxidative stress (Supplementary Fig. S8B; Supplementary Table S6). Notably, mitochondrial localization and protection from oxidative stress required new protein synthesis, since they were blocked by cycloheximide (Supplementary Fig. S8A,B). Furthermore, we found that localization of NKX3.1 to mitochondria, as well as protection from oxidative stress, were not affected by Leptomycin B, an inhibitor of nuclear export, but were blocked by Valinomycin, an inhibitor of mitochondrial import (Supplementary Fig. S9A,B; Supplementary Table S6). These findings suggest that mitochondrial NKX3.1 corresponds to newly synthesized protein imported to mitochondria directly rather than first exported from the nucleus.

Interrogation of the primary sequence of NKX3.1 using the MitoMiner resource tool (53) did not reveal an evident mitochondrial localization domain; however, the lack of a three-dimensional structure for the full length NKX3.1 protein may preclude identification of such domain. Indeed, our findings showing that only full length NKX3.1, but not various truncated proteins, is localized to mitochondria (see Supplementary Fig. S7B), suggests that its tertiary structure is necessary for import.

To gain insight regarding how NKX3.1 is imported to mitochondria, we performed mass spectrometry to identify proteins associated with NKX3.1 in mitochondria following oxidative stress (Fig. 3A,B; Supplementary Dataset 5). Among these, we focused on HSPA9 (Heat Shock Protein Family A Member 9, and also known as Mortalin), a chaperone protein known to import proteins to mitochondria (54) that has various functions in cancer (55-57). Notably, HSPA9 is a non-essential gene, making it feasible to study the consequences of its silencing for NKX3.1 localization to mitochondria and protection from oxidative stress.

Figure 3. HSPA9 imports NKX3.1 to mitochondria to protect from oxidative stress and suppress tumorigenicity.

(A) Strategy to isolate NKX3.1-associated proteins in mitochondria. RWPE1 cells expressing a FLAG-HA-tagged NKX3.1 were treated with paraquat (100μM) for 24 hours followed by biochemical fractionation to isolate mitochondria. NKX3.1-interacting proteins were isolated by immunoprecipitation and identified by mass spectrometry (see Methods). (B) Western blot analysis following co-immunoprecipitation of mitochondria from paraquat-treated RWPE1 cells expressing FLAG-HA-tagged NKX3.1 (or a control vector). NKX3.1 protein complexes were isolated using anti-FLAG followed by Western blot detection with the indicated antibodies. (C) Western blot analysis following co-immunoprecipitation of NKX3.1, T164A, and R52C to detect HSPA9. RWPE1 cells expressing FLAG-HA-tagged proteins (or a control vector) were treated with paraquat (100μM) or vehicle for 24 hours. In panels B and C, input represents 10% of the total protein used for immunoprecipitation; IP, immunoprecipitated proteins. (D, E) RWPE1 cells expressing NKX3.1 (or a control vector) were infected with an shRNA for HSPA9 (shHSPA9) and treated with paraquat (100μM) or vehicle for 24 hours. (D) Western blot analysis of nuclear and mitochondrial fractions. (E) Quantification of mean ROS intensity as detected by DHE production. (F-M) Analysis of LNCaP cells expressing shRNAs for HSPA9 (shHSPA9) and/or NKX3.1 (shNKX3.1) or the control (shControl), as indicated. Cells were treated with paraquat (100μM) or vehicle for 24 hours. (F) Western blot analysis of nuclear and mitochondrial fractions. (G) Confocal images of LNCaP cells co-stained with anti-NKX3.1 (red), anti-HSPA9 (cyan), and anti-ATPB to visualize mitochondria (green); nuclei were visualized by labeling with DAPI (blue). Scale bars represent 25μm. (H and I) Quantification of mean ROS intensity as detected by DHE (H) or MitoSOX (I). (J) Quantification of cellular proliferation as detected by MTT absorbance. (K) Representative images of colony formation assays visualized by staining with crystal violet. (L) Quantification of number of colonies. (M) Quantification of Matrigel invasion assays. Unless otherwise indicated, shown are representative data from 2-3 independent experiments, each done in triplicate (9 independent samples/group). Data are expressed mean ± SD; P values were calculated using a two-sample unpaired Welch t-test. ns, not significant. See also Supplementary Figures S8 and S9, Supplementary Tables S3, S4, and S6, and Dataset 5.

Co-immunoprecipitation assays performed using NKX3.1-expressing or control RWPE1 cells, demonstrated association of NKX3.1 with HSPA9 in mitochondria following oxidative stress (Fig. 3B,C). Notably, NKX3.1(T164A), but not NKX3.1(R52C), interacts with HSPA9 in mitochondria (Fig. 3C), consistent with our findings that NKX3.1(R52C) does not localize to mitochondria. Furthermore, shRNA-mediated silencing of HSPA9 in NKX3.1-expressing RWPE1 cells abolished localization of NKX3.1 to mitochondria and protection from oxidative stress (Fig. 3D,E). We observed similar findings following HSPA9 silencing in LNCaP cells, as evident by Western blot and immunofluorescence analyses (Fig. 3F-H). Moreover, HSPA9 silencing in LNCaP cells impaired the ability of NKX3.1 to protect from oxidative stress-induced acceleration of tumorigenicity in vitro (Fig. 3J-M). These findings demonstrate that import of NKX3.1 to mitochondria by HSPA9 is necessary for protection from ROS and suppression of tumorigenicity.

NKX3.1 regulates the expression of mitochondrial-encoded electron transport chain genes

The mitochondrial genome encodes 37 genes, 13 of which are essential subunits of the electron transport chain (ETC) complexes (Fig. 4A-C), which are necessary for OXPHOS and generate ROS as a byproduct (15-18). Strikingly, in response to oxidative stress, all 13 mitochondrial-encoded ETC genes were up-regulated in NKX3.1-expressing RWPE1 cells, while none were up-regulated in cells expressing either NKX3.1(T164A) or NKX3.1(R52C) (Fig. 4B-D; Supplementary Dataset 2). Similar results were observed in LNCaP cells in which reduced expression of ETC genes in NKX3.1-silenced cells following oxidative stress could be rescued by native NKX3.1, but not NKX3.1(T164A) or NKX3.1(R52C), and also in Nkx3.1^−/− versus Nkx3.1^+/+ mouse prostate tissues (Supplementary Fig. S10A,B; Supplementary Dataset 1). Furthermore, NKX3.1-dependent upregulation of mitochondrial-encoded ETC genes was abrogated by HSPA9 silencing and rescued treatment with MitoQ (Supplementary Fig. S10C-E).

Figure 4. NKX3.1 regulates expression of mitochondrial-encoded electron transport chain genes.

(A) Schematic diagram of OXPHOS complexes of the electron transport chain (ETC) located in the inner mitochondrial membrane. Shown are sites of NAD+ (Complex I) and ATP (Complex V) production as well as the major sources of ROS (Complex I and III). Complexes I, III, and IV act as proton (H⁺) pumps to facilitate electron transport, whereas Complex V moves H⁺ from the intermembrane space to the mitochondrial matrix to convert ADP to ATP. Free electrons lead to ROS accumulation in the form of superoxide (O⁻₂) or hydrogen peroxide (H₂O₂). (B-F) Analyses of RWPE1 cells expressing NKX3.1, T164A, or R52C (or the control vector) treated with vehicle or paraquat (100μM) for 24 hours. (B,C) Heat map representations of nuclear-encoded and/or mitochondrial-encoded (MT-encoded) ETC genes based on RNA sequencing analyses. (B) Differential expression of MT-encoded ETC genes in cells expressing NKX3.1, T164A, or R52C. (C) Differential expression of nuclear-encoded and MT-encoded ETC genes in NKX3.1-expressing cells; MT-encoded genes are shown by arrows. (D) Quantitative real-time PCR analysis showing relative expression of MT-encoded ETC genes. Data are expressed as relative mRNA levels (relative to 18s rRNA expression) showing the mean ± SD; (E-F) NKX3.1 binding to the D-Loop in the mitochondrial genome. (E) Schematic representation of the D-loop, indicating the location of the NKX3.1 consensus DNA binding site (TAAGTA) and the positions of two independent primer sets (primer sets A and B) used for Chromatin immunoprecipitation-quantitative PCR (ChIP-qPCR) (arrows). LSP, light strand promoter position. HSP1, heavy strand promoter 1 position. HSP2, heavy strand promoter 2 position. (F) ChIP-qPCR of NKX3.1 binding to the D-loop in RWPE1 cells expressing exogenous NKX3.1, T164A, or R52C. Cells were treated with paraquat (100μM) or vehicle, and ChIP was done using an antibody against human NKX3.1. Data are expressed as relative enrichment of NKX3.1 binding showing the mean ± SD. Unless otherwise indicated, shown are representative data from 3 independent experiments, each done in triplicate (9 independent samples/group). P values were calculated using a two-sample unpaired Welch t-test. ns, not significant. See also Supplementary Figures S10-S12 and Dataset 2.

In considering mechanisms by which NKX3.1 up-regulates mitochondrial ETC genes, we noticed that the D-Loop, which is the regulatory region in the mitochondrial genome that controls expression of these genes (18), has a perfect NKX3.1 consensus DNA binding sequence, namely TAAGTA (Fig. 4E) (58). Additionally, transcription of ETC genes is necessary for protection from oxidative stress by NKX3.1 since balapiravir, an inhibitor of mitochondrial-specific RNA polymerase (59), abrogated NKX3.1-dependent protection from oxidative stress (Supplementary Fig. S11A,B).

Indeed, using chromatin immunoprecipitation followed by quantitative PCR (ChIP-qPCR), we found that, in response to oxidative stress, NKX3.1 binds specifically to the D-loop in the mitochondrial genome (Fig. 4F; Supplementary S12A). Furthermore, mouse Nkx3.1 bound to the corresponding site in the D-loop of the mouse mitochondrial genome (Supplementary S12B). Interaction of NKX3.1 with the D-loop was dependent on binding to DNA and localization to mitochondria since neither NKX3.1(R52C) nor NKX3.1(T164A) bound to the D-loop, and since binding was abrogated by silencing HSPA9 (Fig. 4F; Supplementary S12C). Therefore, in response to oxidative stress, NKX3.1 regulates transcription of mitochondrial ETC genes, which requires import to mitochondria and binding to a regulatory region of the mitochondrial genome.

NKX3.1 regulates OXPHOS activity and mitochondrial respiration

The mitochondrial-encoded genes regulated by NKX3.1 are essential subunits of the ETC complexes (ETC complexes I-V), which act in concert to regulate OXPHOS functions, beginning with the conversion of NADH to NAD⁺ and culminating with the generation of ATP, and in the process generating mitochondrial ROS (17) (see Fig. 4A). To evaluate whether NKX3.1 affects ETC complex activity, we isolated mitochondria from NKX3.1-expressing or control RWPE1 cells following treatment with paraquat, and measured the activity of each complex as follows: (i) Complex I, NADH ubiquinone oxidoreductase activity; (ii) Complex II, Succinate-coenzyme Q Reductase activity; (iii) Complex III, conversion of oxidized Cytochrome c to its reduced form; (iv) Complex IV, Cytochrome c oxidase activity; and (v) Complex V, ATP synthase activity (see Methods). Activity of all five ETC complexes were significantly reduced in mitochondria isolated from paraquat-treated control RWPE1 cells; however, activity was fully restored in mitochondria isolated from the NKX3.1-expressing cells, but not from cells expressing either NKX3.1(T164A) or NKX3.1(R52C) (Fig. 5A). Restoration of ETC complex activity by NKX3.1 was accompanied by increased expression of the ETC proteins (Fig. 5B); additionally, the requirement for NKX3.1 was overcome by the mitochondrial-specific anti-oxidant, MitoQ (Supplementary Fig. S13A).

Figure 5. NKX3.1 regulates OXPHOS activity and mitochondrial respiration.

(A-D) Analyses of RWPE1 cells expressing NKX3.1, T164A, or R52C treated with paraquat (100μM) or vehicle for 24 hours. (A) Quantification of activity of OXPHOS complexes I, II, III, IV and V from mitochondria isolated from paraquat-treated cells (n=5 independent samples/group). (B) Western blot analyses of nuclear and total mitochondrial fractions to assess expression levels of the mitochondrial-encoded (mt) OXPHOS complex proteins as indicated. (C) NADH/NAD⁺ ratio. (D) Relative ATP levels. (E-H) Seahorse analyses of mitochondrial respiration and glycolysis in NKX3.1-expressing (or control) RWPE1 and NKX3.1-silenced (or control) LNCaP cells. (E and F) Oxygen consumption rate (OCR) analyses. The rates of basal-, ATP-linked, maximal, and reserve respiration were quantified by normalization of OCR level to the total protein optical density (OD) values. (G and H) Extracellular acidification rate (ECAR) analyses. The rates of glycolysis, glycolytic capacity and reserve were quantified by normalization of ECAR level to the total protein OD values. Unless otherwise indicated, shown are data from 3 independent experiments, each done in triplicate (9 independent samples/group); P values were calculated using two-sample unpaired Welch t-test. ns, not significant. See also Supplementary Figure S13.

To further investigate the consequences of NKX3.1 for ETC activity, we measured the ratio of NADH and NAD⁺ as well as the levels of ATP, which correspond to the first and last steps of the electron transport chain, respectively (see Fig. 4A), although not exclusively produced via OXPHOS. Paraquat-treatment of control RWPE1 cells resulted in an increase in the NADH/NAD⁺ ratio and a corresponding reduction in ATP levels relative to vehicle-treated cells, whereas both NADH/NAD⁺ and ATP levels were restored in NKX3.1-expressing RWPE1 cells, but not cells expressing NKX3.1(T164A) or NKX3.1(R52C) (Fig. 5C,D). Similarly, in mouse, paraquat treatment resulted in an increased NADH/NAD⁺ ratio and reduced ATP levels in Nkx3.1^−/− relative to Nkx3.1^+/+ prostate (Supplementary Fig. S13B,C).

Furthermore, paraquat treatment of control RWPE1 cells or NKX3.1-silenced LNCaP cells resulted in a significant reduction in OXPHOS (oxygen consumption rate, OCR) and a corresponding increase in glycolytic metabolism (extracellular acidification rate, ECAR), which were restored in the corresponding NKX3.1-expressing cells (Fig. 5E-H). This was accompanied by increased production of L-lactate in the control RWPE1 cells, which was fully restored to physiological levels in NKX3.1-expressing cells, but not in those expressing NKX3.1(T164A) or NKX3.1(R52C), and rescued by MitoQ (Supplementary Fig. S13D-F). Therefore, NKX3.1 restores energy balance in conditions of oxidative stress, thereby promoting mitochondrial homeostasis.

Association of NKX3.1 expression levels with mitochondrial function and clinical outcome in human prostate cancer

To investigate the relevance of these findings for human prostate cancer, we established organotypic cultures from freshly-collected prostate tumor specimens obtained directly from the surgical suite (Fig. 6A). We cultured tissues for up to 5 days, during which time they retained the histological features of the corresponding primary tumors, including expression of NKX3.1 (Fig. 6B). Notably, while NKX3.1 was localized primarily to nuclei in standard culture conditions, paraquat treatment resulted in increased non-nuclear protein (Fig. 6B); confocal imaging showed co-localization of non-nuclear NKX3.1 with mitochondrial markers (Fig. 6C). Localization of NKX3.1 to mitochondria following oxidative stress was accompanied by binding to the D-loop of the mitochondrial genome (Fig. 6D), indicating that the mitochondrial NKX3.1 is active in the organotypic cultures.

Figure 6. Association of NKX3.1 expression levels with mitochondrial function and clinical outcome in human prostate cancer.

(A-H) Analyses of primary human organotypic cultures. (A) Strategy. Primary human prostate cancer tissues were obtained directly from surgery and cultured in vitro in media alone (vehicle) or media containing paraquat (100μM) for 24 hours. (B) Representative images showing histology (H&E) and NKX3.1 immunostaining. Scale bars represent 25μm. (C) Representative confocal images co-stained for NKX3.1 (red) and ATPB (green); nuclei were visualized with DAPI (blue). Scale bars represent 25μm (left and center) or 200μm (right). (D) ChIP-qPCR of NKX3.1 binding to the D-loop of the mitochondrial genome in human prostate organotypic cultures. ChIP was done using an antibody that recognizes human NKX3.1. Data are expressed as relative enrichment of NKX3.1 binding showing the mean ± SD. (E-H) Analyses of human prostate organotypic cultures having “high” versus “low” levels of NKX3.1 expression. (E) Heat map showing NKX3.1 expression levels determined by RT-qPCR analyses. (F) RT-qPCR analysis of mitochondrial-encoded ETC genes in paraquat- versus vehicle-treated organotypic cultures. Data are expressed as relative mRNA levels (relative to 18s rRNA expression) showing the mean ± SD. (G) NADH/NAD⁺ ratio. (H) ATP levels. Panels F-H shows representative data from 3 independent experiments, each done in triplicate (9 independent samples/group). P values were calculated using two-sample unpaired Welch t-test. ns, not significant. (I) Biological pathway-based GSEA using a gene signature comparing “high” versus “low” NKX3.1 mRNA expression from the Taylor cohort to query pathways from the C2 pathway collection (see Methods). “High” corresponds to the top 25% of patients with highest levels of NKX3.1 expression (n=32) and “Low” corresponds to the top 25% of patients with the lowest levels of NKX3.1 expression (n=32). Normalized Enrichment Score (NES) and P values were calculated based on 1000 permutations. (J) Kaplan-Meier survival analysis showing association of NKX3.1 and mitochondrial-encoded ETC (mito) expression levels with biochemical recurrence (BCR) estimated survival probability based on the TCGA cohort. Analyses compare patients with ‘high’ or ‘low’ expression levels of NKX3.1 and “high” or “low” expression levels of the combined 13 mitochondrial ETC genes (mito); P values were estimated using a log-rank test. See also Supplementary Figure S14, Supplementary Tables S7 and S8, and Dataset 6.

We and others have shown that expression of NKX3.1 in prostate cancer is heterogenous both at the mRNA and protein levels (34,36,37,44). In particular, human prostate cancer patients can be segregated based on having low- or high-levels of NKX3.1, which is associated with their differing response to a chemo-preventive agent (36). We now find that the organotypic cultures can also be separated based on having low or high NKX3.1 mRNA expression levels (Fig. 6E, Supplementary Table S7), thereby making it feasible to evaluate response to oxidative stress relative to NKX3.1 expression levels. We found that paraquat treatment resulted in up-regulation of mitochondrial ETC gene expression in organotypic cultures having high versus low levels of NKX3.1 (Fig. 6F); correspondingly, we observed an increased NADH/NAD⁺ ratio and reduced ATP levels in the paraquat-treated cultures with low versus high levels of NKX3.1 (Fig. 6G,H).

To identify biological pathways enriched in tumors with high versus low levels of NKX3.1, we queried expression profiles from a cohort of primary prostate tumors (Taylor cohort, n=131; Supplementary Table S8) (60). Enriched pathways included those associated with oxidative stress, metabolism, and mitochondrial homeostasis (Fig. 6I; Supplementary Dataset 6), several of which are conserved between mouse and human prostate (see Supplementary Fig. S1). To examine the association of expression levels of NKX3.1 and those of mitochondrial ETC genes with clinical outcome, we queried expression profiles from a cohort of primary tumors from The Cancer Genome Atlas (TCGA) (n=426; Supplementary Table S8) (61), which has available expression data for nuclear as well as mitochondrial genes (62), including the 13 mitochondrial ETC genes regulated by NKX3.1 (see Fig. 4). We grouped the patients based on having low (the 25% lowest, n=107) versus high (the 25% highest, n=107) expression levels of NKX3.1 or combined expression levels of the 13 mitochondrial ETC genes, such that we compared 4 patient groups with high-high (n=20), high-low (n=68), low-high (n=18) or low-low (n=62) expression levels of NKX3.1 and ETC genes. Patients with low expression of NKX3.1 and ETC genes had worse clinical outcome than any of the other combinations (Log-rank p-value=5.8X10⁻⁴, Fig. 6J), which underscores the prognostic relevance of NKX3.1 and mitochondrial ETC expression for prostate carcinogenesis. Interestingly, NKX3.1 expression levels were also correlated with those of HSPA9 in this cohort (p-value=1.6X10⁻⁵, Supplementary Fig. S14A).

Association of non-nuclear NKX3.1 with clinical outcome in human prostate cancer

To investigate whether NKX3.1 protein expression levels and sub-cellular localization are associated with clinical outcome we used two independent human prostate cancer tissue microarrays (TMA-1 and TMA-2). TMA-1 is comprised of low Gleason primary tumors (Gleason 6 and 7) with a limited clinical follow-up period (n=194, Supplementary Table S8), while TMA-2 is comprised of primary tumors with the full range of Gleason scores (Gleason 5 to 9) with long-term clinical follow-up data (n=118, Supplementary Table S8 (30)). We performed immunostaining for NKX3.1 to evaluate staining intensity (high, low, or negative) for nuclear and non-nuclear NKX3.1 and to correlate staining levels with Gleason score, PSA levels, and clinical outcome (Fig. 7A-G; Supplementary Fig. S14B-E). Immunostaining for NKX3.1 revealed both nuclear and non-nuclear expression (Fig. 7B), while confocal imaging showed co-localization of non-nuclear NKX3.1 with mitochondrial markers (Fig. 7C), consistent with its localization to mitochondria. Expression of NKX3.1 was positively correlated with HSPA9, and negatively with 4HNE, a marker of lipid oxidation (Fig. 7D).

Figure 7. Association of non-nuclear NKX3.1 protein with clinical outcome in human prostate cancer.

(A) Strategy. NKX3.1 protein expression levels were examined on two independent TMAs. TMA1 is comprised of 194 patients with Gleason score 6 or 7 tumors; TMA2 is comprised of 118 patients with Gleason score 5-9 tumors. Immunostaining of NKX3.1 was graded based on nuclear or non-nuclear expression and corresponding to “high”, “low”, or “negative” expression. (B) Representative cases based on TMA2 showing nuclear or non-nuclear NKX3.1 immunostaining and examples of “high”, “low”, or “negative” expression. Scale bars represent 50μm. (C) Representative confocal images of human prostate tissues co-stained for NKX3.1 (red) and ATPB (green); nuclei were visualized with DAPI (blue). Scale bars represent 25μm. (D) Representative human prostate tumors showing examples of “high” or “low” NKX3.1. Adjacent sections were stained for HSPA9 and 4HNE (marker of oxidative damage). (E) Analyses of TMA2, showing the association of “high” or “low/no” expression levels of total, nuclear, or non-nuclear NKX3.1 protein relative to pre-operative PSA levels (total n of patients = 82). P values were calculated using two-sample unpaired Welch t-test. ns, not significant. (F, G) Kaplan-Meier survival analyses showing association of total, nuclear, or non-nuclear NKX3.1 protein expression levels with biochemical recurrence (BCR)-free estimated survival probability for TMA1 (n=112) (Panel F) and overall survival probability for TMA2 (n=90) (Panel G); P values were estimated using a log-rank test. See also Supplementary Figure S14 and Supplementary Table S8.

We separated patients based on their having high or low levels of NKX3.1 considering total protein (e.g., both nuclear and non-nuclear), or only nuclear or only non-nuclear protein (Fig. 7A,B). Although expression levels of NKX3.1 varied across Gleason scores, consistent with previous studies (e.g., (34,37)), we found that patients with higher Gleason scores tended to have low levels of non-nuclear NKX3.1, which was not the case when we considered total or only nuclear NKX3.1 protein (Supplementary Fig. S14B). Furthermore, patients with low levels of non-nuclear NKX3.1 had significantly higher PSA levels, whereas no difference in PSA levels was observed in patients based on total or nuclear NKX3.1 protein (Fig. 7E; Supplementary Fig. S14C).

Most notably, we observed a significant association of NKX3.1 protein expression levels and sub-cellular localization with clinical outcome for both TMA-1 and TMA-2 (Fig. 7F,G; Supplementary Fig. S14D,E). In particular, Kaplan Meier survival analyses revealed that reduced expression of NKX3.1 is associated with adverse disease outcome when estimated based on biochemical recurrence for TMA-1 (Log-rank p value = 0.0232) or overall survival for TMA-2 (Log-rank p value = 0.0063) (Fig. 7F,G). Furthermore, a multivariable Cox proportional hazards model, adjusted for Gleason score only or all clinical variables showed that the relationship of NKX3.1 expression to outcome is independent of these variables (hazard ratio HR=4.7, P = 0.019 and HR=7.4, P=0.032, respectively) (Supplementary Fig. S14D). Most notably, the association of NKX3.1 protein levels with outcome was primarily accounted for by reduced expression of non-nuclear NKX3.1 protein (Fig. 7F,G; Supplementary Fig. S14E). In particular, low levels of nonnuclear, but not the nuclear, NKX3.1 protein were significantly associated with adverse outcome estimated based on biochemical recurrence for TMA-1 (Log-rank p value = 0.0170, Fig. 7F; Log-rank p value = 0.027, Supplementary Fig. S14E), and overall survival for TMA-2 (Log-rank p value = 0.0063, Fig. 7G; Log-rank p value = 0.0079, Supplementary Fig. S14E). These findings demonstrate the significance of non-nuclear NKX3.1 for clinical outcome in prostate cancer.

Discussion

Mitochondria provide a front line of defense for protecting cells from the adverse consequences of oxidative stress in normal and abnormal contexts. Mitochondrial homeostasis is controlled by a series of protein complexes that comprise the electron transport chain (ETC), which are comprised primarily of nuclear-encoded genes, although a few essential ETC proteins are encoded by the mitochondrial genome. We now show that, in response to oxidative stress, the nuclear transcriptional factor NKX3.1 is recruited to mitochondria where it regulates expression of these mitochondrial-encoded ETC genes, thereby promoting mitochondrial homeostasis and suppressing cancer initiation.

We propose a model in which NKX3.1 localization to mitochondria provides protection from the adverse consequences of oxidative stress, which contributes to its ability to suppress cancer initiation (Supplementary Fig. S15). Conversely, in scenarios in which NKX3.1 localization to mitochondria is low or impaired, loss of such protection leads to aberrant mitochondrial integrity and accelerated cancer initiation (Supplementary Fig. S15). Thus, our findings elucidate a hitherto unappreciated role of NKX3.1 in mitochondria for suppression of prostate cancer initiation, and provide new insights regarding how communication between the nucleus and mitochondria might sustain ETC function and ROS levels in response to oxidative stress.

Indeed, relatively few nuclear transcription factors have been shown to function in mitochondria, and even fewer to directly regulate the expression of mitochondrial genes (63,64). Notable among these is the androgen receptor (AR), which, like NKX3.1, is a key regulator of prostate differentiation and cancer. However, in contrast to NKX3.1, which increases mitochondrial ETC gene expression and to restores OXPHOS function, AR negatively regulates expression of ETC complexes, which is associated with mitochondria dysfunction (65). This raises the interesting possibility that NKX3.1 and AR counterbalance each other in regulating mitochondrial homeostasis in response to oxidative stress. Indeed, prostate epithelial cells, which express both NKX3.1 and AR (2), are highly susceptible to oxidative stress that arises as a consequence of cellular senescence, inflammation, DNA damage, aging or other factors (7,11).

Unexpectedly, mitochondrial NKX3.1 does not originate from the nucleus; rather, in response to oxidative stress newly synthesized NKX3.1 is imported to mitochondria by HSPA9. Nonetheless, the transcriptional properties of mitochondrial NKX3.1 appear similar to those of nuclear NKX3.1, since transcription of mitochondrial ETC genes is mediated by binding via the homeodomain to an NKX3.1 consensus DNA binding site in the mitochondrial genome. The significance of these biochemical data is underscored by our finding that patients with low expression levels of NKX3.1 and mitochondrial ETC genes have less favorable clinical outcome, compared with their counterparts that have high expression levels of NKX3.1 and/or ETC genes.

Interestingly, regulation of mitochondrial ETC gene expression by NKX3.1, which is a prostate specific homeoprotein, occurs via binding to an essential regulatory region of the mitochondrial genome, called the D-loop. The occurrence of a tissue-specific NK-class homeodomain consensus binding site within a ubiquitous and highly conserved region of the mitochondrial genome raises the question as to whether similar regulation occurs in other tissue-specific contexts. Indeed, other tissue-specific NK-class homeoproteins have been shown to control ROS levels and effect mitochondrial function, as well as to exhibit non-nuclear localization (66-68), although they have not been reported to be localized to mitochondria.

While association of NKX3.1 expression status with prostate cancer progression has been well studied (26,34,36,37,43), most previous studies have focused on nuclear NKX3.1. Our current findings unveil a new dimension of NKX3.1 regulation since we demonstrate association of non-nuclear NKX3.1 with cancer outcome. We propose that analyses of NKX3.1 expression status combined with analyses of its sub-cellular localization may inform precision prevention monitoring, particularly for men on active surveillance.

Furthermore, NKX3.1 is associated with hereditary prostate cancer and its loss of expression has been linked to increased risk of aggressive prostate cancer, especially among African American men (35). We find that two polymorphisms of NKX3.1, each of which is associated with increased risk of aggressive prostate cancer (51,52), encode proteins that are incapable of functioning in mitochondria to protect against oxidative stress or preserve mitochondrial function. These data suggest a mechanism directly linking NKX3.1 status to prostate cancer patients that are at higher risk, opening new avenues for genetic risk assessment in prostate cancer.

Methods

Analysis of Nkx3.1 germline mutant mice

All experiments using animals were performed according to protocols approved by the Institutional Animal Care and Use Committee. Nkx3.1^+/+ and Nkx3.1^−/− mice (27) were administered paraquat dichloride/methyl viologen dichloride hydrate (10 mg/kg/day, Sigma-Aldrich, St. Louis, MO, USA) or vehicle for 1 to 9 months. Histopathological analysis and immunohistochemical staining were done as described (39). A list of all antibodies used in this study is provided in Supplementary Table S9.

Analysis of human prostate cells

RWPE1 and LNCAP prostate cancer cells were obtained from the American Type Culture Collection (ATCC); BPH1 cells were obtained from EMD Millipore (Burlington, MA, USA). Cells were maintained under mycoplasma-free conditions (30-1012K, ATCC) and only low passage cells were used. All procedures for lentiviral studies were approved by the Office of Environmental Health and Safety. Lentiviruses were generated using second generation packaging vectors (psPAX2 and pMD2.G, Addgene, Cambridge, MA, USA). Procedures for their preparation and transduction were as described (39). A list of all primers and shRNA used in this study is provided in Supplementary Table S10.

To induce oxidative stress, cells were treated with 100μM paraquat dichloride/methyl viologen dichloride hydrate (PQ) (Sigma-Aldrich, St. Louis, MO, USA) or 50μM MitoParaquat (MitoPQ) (Cayman Chemical, Ann Arbor, MI, USA). Measurement of reactive oxygen species (ROS) levels was done using: (i) 10μM DHE (Invitrogen, Carlsbad, CA, USA); (ii) 10μM CM-H2DCFDA (Invitrogen, Carlsbad, CA, USA); (iii) 5μM MitoPY1 (Tocris Bioscience, Minneapolis, MN, USA); or (iv) 10μM MitoSOX (Invitrogen, Carlsbad, CA, USA). FACS analysis was done using excitation/emission spectra of: (i) A566/A616 for DHE; (ii) A595/A519 for CM-H2DCFDA; or (iii) A595/519 for MitoPY1; or (iv) A596/A576 for MitoSOX. Proliferation, colony formation and invasion assays were performed as described (28).

Gene expression profiling analysis

RNA sequencing was performed as previously described (39) on anterior prostate of Nkx3.1^+/+ or Nkx3.1^−/− mice, or NKX3.1-expressing (or control) human RWPE1 cells. Differential expression signatures were ranked by t-values from a two-tailed two-sample Welch t-test. Pathway enrichment was done using GSEA to query pathways collected from the C2 database. Heatmap depiction of the differential expression of ETC genes was done using https://www.wikipathways.org/ (RRID:SCR_002134).

Sub-cellular localization analyses

Electron microscopy (EM) was performed as described (69). Immuno-electron microscopy was done on cells fixed in 4% paraformaldehyde plus 1% glutaraldehyde (Electron Microscopy Sciences, Fort Washington, PA, USA) and embedded in Lowicryl HM20 resin (Electron Microscopy Sciences, Fort Washington, PA, USA). Biochemical fractionation of organelles was done as described (70). Measurement of mitochondrial membrane potential using TMRE was done following the manufacturer’s instructions (Abcam, Cambridge, MA, USA). Measurement of mitochondrial mass was performed using MitoTracker Red CMXRos (M7512, Invitrogen, Carlsbad, CA, USA) or MitoView Green (70054, Biotium, Fremont, CA, USA) as per the manufacturer’s instructions.

DNA binding studies

Gel retardation assays were done as described (39), using protein extracts from nuclear or mitochondrial cell fractions. Chromatin immunoprecipitation was done on mitochondria genomic DNA from cells or tissue (as above) followed by real time PCR (ChIP-qPCR analysis).

Cellular metabolism assays

For measurement of ETC complex activity, the Mitochondria Isolation Kit (ab110169, Abcam, Cambridge, MA, USA) was used to isolate mitochondria, and complex I-V enzymatic activity were measured by using assay kits for complex I (ab109903, Abcam, Cambridge, MA, USA), complex II (700940, Cayman Chemical, Ann Arbor, MI, USA), complex II+III (ab109905, Abcam, Cambridge, MA, USA), complex IV (ab109906, Abcam, Cambridge, MA, USA), and complex V (ab109907, Abcam, Cambridge, MA, USA), following the manufacturer’s instructions. Oxygen consumption rate (OCR) and extracellular acidification rate (ECAR) were quantified using a Seahorse XF96 (Agilent Technologies, Santa Cruz, CA, USA). Quantitative analysis of ATP, NADH/NAD⁺ ratio, and lactate levels were performed according to manufacturer’s recommendations (ATP assay kit, Sigma-Aldrich, St. Louis, MO, USA; NADH/NAD quantitation kit, Sigma Aldrich, St. Louis, MO, USA; L-Lactate Assay kit, Abcam, Cambridge, MA, USA).

Analysis of human prostate cancer

All studies using anonymized human prostate specimens were performed according to the Declaration of Helsinki protocol and protocols approved by the Human Research Protection Office and Institutional Review Board at Columbia University Irving Medical Center. All studies were done with written informed consent from the patients.

Statistical Analyses

For in vivo studies, the number of mice needed to achieve statistical significance was determined using standard power analysis. All cell culture analyses were done using a minimum of three biological replicates, each done in triplicate. Statistical analyses were performed using two-sample unpaired Welch t-test, Chi-squared (x²) test, Fisher’s exact test or Spearman correlation test as indicated in each figure legend. For the box plots, boxes indicate the 25th–75th percentiles, center-lines show the median, and whiskers show the minimum–maximum values. Statistical analysis was performed using GraphPad Prism software (Version 8.4.3, RRID:SCR_002798) and R-studio (0.99.902, R v4.0.2). Cox proportional hazards and survival analysis was performed using survival and survminer packages in R-studio.

Data Availability

The RNA sequencing expression profiling data have been deposited in the Gene Expression Omnibus (GEO) database (https://www.ncbi.nlm.nih.gov/geo/, RRID:SCR_005012) with the accession code GSE115338.