Next Article in Journal
Recent Progress of Stem Cell Therapy in Cancer Treatment: Molecular Mechanisms and Potential Applications
Previous Article in Journal
Cross-Presenting XCR1+ Dendritic Cells as Targets for Cancer Immunotherapy
Open AccessReview

Exosomes are the Driving Force in Preparing the Soil for the Metastatic Seeds: Lessons from the Prostate Cancer

Laboratory of Molecular Cell Biology, Department of Zoology, Faculty of Science, Assiut University, Assiut 71515, Egypt
Department of Pharmaceutical Sciences, Rangel College of Pharmacy, Texas A&M Health Science Center, College Station, TX 77843, USA
Department of Surgery, School of Medicine, Tulane University, New Orleans, LA 70112, USA
Author to whom correspondence should be addressed.
Cells 2020, 9(3), 564;
Received: 21 January 2020 / Revised: 17 February 2020 / Accepted: 20 February 2020 / Published: 28 February 2020
(This article belongs to the Section Cellular Pathology)


Exosomes are nano-membrane vesicles that various cell types secrete during physiological and pathophysiological conditions. By shuttling bioactive molecules such as nucleic acids, proteins, and lipids to target cells, exosomes serve as key regulators for multiple cellular processes, including cancer metastasis. Recently, microvesicles have emerged as a challenge in the treatment of prostate cancer (PCa), encountered either when the number of vesicles increases or when the vesicles move into circulation, potentially with an ability to induce drug resistance, angiogenesis, and metastasis. Notably, the exosomal cargo can induce the desmoplastic response of PCa-associated cells in a tumor microenvironment (TME) to promote PCa metastasis. However, the crosstalk between PCa-derived exosomes and the TME remains only partially understood. In this review, we provide new insights into the metabolic and molecular signatures of PCa-associated exosomes in reprogramming the TME, and the subsequent promotion of aggressive phenotypes of PCa cells. Elucidating the molecular mechanisms of TME reprogramming by exosomes draws more practical and universal conclusions for the development of new therapeutic interventions when considering TME in the treatment of PCa patients.
Keywords: castrate resistant prostate cancer; tumor microenvironment; stromal cells; exosomal cargo castrate resistant prostate cancer; tumor microenvironment; stromal cells; exosomal cargo

1. Introduction

Prostate cancer (PCa) is the most common adenocarcinoma in American and European men, after skin cancer [1,2]. As estimated by the American Cancer Society, approximately 174,650 new cases and 31,620 deaths from PCa were predicted annually in the United States as of 2019 [3]. In early-stage PCa, the cancer cells remain sensitive to androgens; therefore, androgen deprivation therapy is the most effective treatment typically offered to these PCa patients [4]. Over time, however, the cancer cells become androgen insensitive, and chemotherapy agents, such as docetaxel, are one clinical option to treat androgen-independent and metastatic castrate-resistant PCa (mCRPC), a stage at which the clinical outcomes of the PCa patient are inferior [5,6]. CRPC is characterized by progression, despite the patient living with castrate levels of testosterone < 0.5 ng/mL [7]. The mechanisms proposed to illustrate this phenomenon include androgen receptor (AR) gene mutation, AR splice variant expression, AR overexpression, an increase in the expression of the activator transcription factors, and up-regulation of the androgen synthesis enzymes, such as CYP17 [8,9,10,11,12,13]. Therefore, although castration levels of the androgen are present in CRPC, the AR signaling pathway remains active. Understanding these pathways will help in the development of new targeting agents to block the AR pathway. These targeting agents include abiraterone, which blocks CYP17A1, a microsomal enzyme involved in two critical steps of testosterone biosynthesis [14,15,16], whereas Orteronel (TAK-700) and Galeterone (TOK-001) work as AR blockers by inhibiting CYP17 [17,18,19]. Common AR antagonists include Enzalutamide (MDV 3100), ARN-509, and ODM-201, which are introduced as therapeutic agents against mCRPC [20,21]. Many of the novel cytotoxic chemotherapeutic agents developed in recent years, such as docetaxel and cabazitaxel, are associated with an increase in the overall survival of mCRPC patient from 9–18 months to > 30 months [22,23,24,25]. PCa expresses tumor-associated antigens, which make cancer cells a target for vaccines [26]. Immunotherapy is an attractive therapeutic approach for treating PCa. For example, Sipuleucel-T is a cell-based immunotherapy and PROSTVAC-VF is a recombinant vaccine that consists of two vectors encoding prostate-specific antigen (PSA) and three immune co-stimulatory agents [27]. Although the mCRPC treatment landscape has developed significantly in the last decade, nonetheless mCRPC patients continue to face a variety of therapeutic challenges that require additional research attention. Today, the impact of the tumor microenvironment (TME) in prostate cancer development and metastasis is commonly highlighted throughout the related literature.

2. The Soil/Seed Analogy: Tumor Microenvironment (TME) and Tumor Cells

Analysis of the TME has been out of reach for many decades, with studies in this area only recently gaining significant momentum in cancer research. The relation between cancer cells and their TME is quite similar to the “seeds and soil” relationship, which explains the tactical role of the TME in cancer evolution and progression as a result of the stimulatory or inhibitory signals that the TME provides [28]. The TME includes the diverse cells in the vicinity of the tumor, such as fibroblast, endothelial, immune, fat, neural, epithelial, and mesenchymal stem cells [29], as well as the soluble and insoluble factors, extracellular matrix and exosomes [30]. Although multiple studies have focused on the modulating role of soluble factors on the TME, new evidence for the potential role of exosomes in altering the TME and promoting aggressive tumor behavior has now been documented [31].

3. Tumor-Associated Exosomes Modulate the TME and Prepare the Metastatic Niche

3.1. Exosomes, Biogenesis, Trafficking, Uptake and Exosomal Cargo

Cells communicate with each other by releasing different types of extracellular vehicles (EVs), such as exosomes, which are cup-shaped bi-layered membrane nanovesicles (30–120 nm in diameter), into their local microenvironment and the circulatory system. EVs are small, double-membrane bodies released by normal and abnormal cells and are classified into three main types based on the size of vesicles. The typical size of EVs ranges from 100 nm to 1µm, exosomes from 30 to120 nm, and apoptotic bodies from 500 nm to 2 µm in diameter [32]. Exosomes are intraluminal vesicles that are derived from multivesicular bodies through a process of endosome ripening, in which the vesicles either fuse with lysosomes to degrade their cargo or fuse with the plasma membrane to release exosomes into the extracellular matrix [33,34,35]. The number and composition of exosomes depend on the physiological cellular activities in which exosomes are involved in. Exosomes can be characterized by a set of exosome markers expressed on the outer membrane of these vesicles or enclosed in their cargo. The most common protein contents of exosomes are tetraspanins (CD63, CD9, CD81, and CD82), ESCRT-I associated protein (TSG101), lysosome-associated membrane glycoproteins (LAMP-1 and 2B), MVB-associated protein (Alix-1), heat shock proteins (HSP60, 70, and 90), adhesion molecules (CD54 and CD11b), major histocompatibility molecules (MHC-I and II), Ras-related proteins (Rabs), and membrane-binding proteins (annexins) [36]. In addition to proteins, exosomes carry in their cargos RNAs (microRNAs, mRNAs, and lncRNAs), DNAs, and lipids, which have been previously reported [37].
Several mechanisms regulate exosomal trafficking and release. Kirsten Rat Sarcoma (RAS)-associated binding (Rab) proteins mediate exosomal trafficking and their release to the extracellular matrix [38]. Interestingly, cancer cells overexpress different EV-associated biogenesis machinery, such as components of the Endosomal Sorting Complex Required for Transport (ESCRT), syntenin, heparanase, YKT6, amplifying Rho/ROCK, EGFRvIII, H-RASV12, and proto-oncogene Src signaling, which subsequently causes the release of a significantly higher quantity of exosomes than normal cells would release [39,40,41,42,43,44,45,46,47,48]. Hypoxic conditions and a low-pH TME can also positively impact the release and uptake of exosomes by cancer cells [49,50,51,52]. In addition, p53 plays a significant role in enhancing exosome secretion, although its mechanism of release remains unknown [53]. Another factor is the ceramides, which can induce exosomal budding into the multivesicular endosomes and inhibit the neutral sphingomyelinase 2 (nSMase2) enzyme, which is a rate limiting enzyme in ceramide biosynthesis, and thereby suppresses exosomal release [54,55,56].
Essentially, the composition of the exosomes determines their autocrine, paracrine, and endocrine functions. The protein cargo of exosomes depends mainly on the ubiquitination process, the plasma membrane anchor tags provided by myristoylation, prenylation, or palmitoylation, and the transmembrane glycoprotein CD43 [57,58]. Even so, it should not be assumed that exosomal RNAs are randomly loaded; in contrast, the process is tightly regulated by specific shuttle events, such as SUMOylated heterogeneous nuclear ribonucleoprotein A2B1 (hnRNPA2B1), which specifically binds to microRNAs (miRNAs or miRs) that contain the ‘shuttling’ motif GGAG, thereby resulting in their upload into exosomes [59]. In addition, it has been suggested that AGO2, a protein accompanied with the RNA-induced silenced complex (RISC), regulates the loading of miRNAs into exosomes [60]. The overexpression of miRNAs and low expression of their target mRNAs are key factors for the loading of these miRNA sequences into exosomes [61]. Furthermore, exosomes protect their cargo against enzymatic degradation through trafficking in the circulatory or extracellular environment [38]. Interestingly, exosomes mirror the metabolic status of the cells of origin, which reflects the emerging role of exosomes as a fingerprint in the diagnosis of many diseases, including PCa [62,63,64,65]. As established by multi-omic studies, tumor-derived exosomes shuttle many bioactive molecules, such as proteins, nucleic acids, and lipid molecules, to and from the TME, which directly exerts phenotypic changes in recipient cells and promotes cancer progression [66]. In the same context, exosomes can communicate between two different cells through 1) transfer of bioactive molecules in their cargo to activate/suppress signaling pathways in target cells, 2) receptors shuttling between donor and recipient cells to alter cellular activities, 3) transfer of fully functional proteins to perform specific functions in target cells, and 4) providing new genetic information to recipient cells to gain new traits [67].
The clinical relevance of exosomes in cancer progression is well established, and differences have been observed in the exosomal cargo of cancerous and normal cells. As such, exosomes have a significant role in the early diagnosis of cancer [68] In PCa patients, several drawbacks remain with regard to the clinical utility of prostate specific antigen (PSA) and carbohydrate antigens as diagnostic markers [69]. However, biopsy is a decisive method of diagnosis, and novel early diagnostic biomarkers are required for clinical applications. The exosomes isolated from PCa blood, urine or saliva can be used as predictive biomarkers. It was reported that PCa patient-derived exosomes shuttled Epidermal Growth Factor Receptor (EGFR) which is overexpressed in PCa tissues at advanced stages. Therefore, enriched EGFR in blood can be used as a noninvasive biomarker that reflects the state of the disease in PCa patients [70]. Khan et al. reported that survivin, an oncoprotein associated with chemoresistance, is overexpressed in patient-derived exosomes and acts as a diagnostic and prognostic marker of PCa [71]. More interestingly, exosomes isolated from serum of African American men with PCa are a wealthy source of biomarkers for early detection and monitoring PCa patients [72]. In addition, exosomes isolated from urine can be utilized as a sentinel to monitor PCa stages and reduce the number of unnecessary biopsies [73].
A previous report indicated that miR-1290 and miR-375 in plasma have a potential role in the prediction of CRPC patients [74]. Exosomal miR-34a predicts docetaxel-treatment failure; that is, this miR contributes to the sensitivity of PCa cells to docetaxel through the downregulation of Bcl-2 [75]. Like miRNAs, exosomal proteins such as annexin A2, calsyntenin 1, fatty acid synthesis, filamin C, folate hydrolase-1, and growth differentiation factor 15 (GDF15) are specific for PCa diagnosis [67]. In addition, exosomal survivin is considered a promising biomarker for the early detection of PCa [71]. Different studies have elucidated the role of exosomes in chemo-resistance [76,77], radio-resistance [78], and immune-resistance [79,80]. It has been suggested that to override the negative effect of exosomes in cancer treatment, exosomes could be depleted from the blood of cancer patients using a hemodialysis-like technique [81]. In addition, exosomes could offer advantages in cancer therapy, such as being used as vehicles for drug delivery. The targeting of cancer cells by specific antibodies or ligands of highly expressed membrane receptors raised against cancer-associated exosomes constitutes an additional example of their therapeutic applications. Exosomes can be used to shuttle miRNAs, siRNA, and anticancer drugs directly to the targeted tumor cells. For example, exosomes loaded with siRNA are able to specifically target neuronal cells in murine brain [82]. In another study, exosomes were successfully delivered Let-7a miRNA to breast cancer cells in a xenograft mouse model [83].
Defining the role of exosomes in cancer biology has gained significant momentum in recent years, although relatively few studies have focused on the potential role of exosomal cargos in the recruitment of PCa neighboring cells to support tumor expansion and metastasis in the “seeds, soil and fertilizer” model. In this review, we have summarized the previously conducted studies on cancer-associated exosomes to better understand the role of bioactive molecules transported by exosomes as they bridge the crosstalk between tumor cells and their TME. The ultimate goal is to offer alternative or complementary therapeutic strategies that account for the TME as a vital component of the tumor architecture.

3.2. Singing Together: PCa Cell-Cell Interaction via Exosomal Signals

Cancer cells communicate with each other via exosomal cargo, which may include signaling complexes, receptors, functional proteins, or genetic information that regulates the signal networks involved in cancer growth and aggressiveness [84,85]. The two-way talk between sister tumor cells ominously affects the advancement of cancer and determines the response of cells to treatment options [86]. Several studies show that exosomes released from cancer cells promote tumor cell proliferation. Soekmadji et al. reported that the increase in androgen-deprived LNCaP cell proliferation was correlated with the release of CD9-positive exosomes [87]. Another study showed that LNCaP- and DU-145-derived exosomes can induce cell proliferation, epithelial-mesenchymal transition (EMT), migration, and IL-8 secretion while decreasing apoptosis in PCa cells [88,89,90]. In addition, hypoxic PCa cell-derived exosomes convey information that can be directly involved in the invasiveness and motility of dormant PCa cells [91]. Moreover, the exosomes released by different PCa cells confer a number of functional proteins to recipient cells, which do not originally express these proteins. It has also been shown that integrins are shuttled by exosomes isolated from PC-3 (αυβ6 and αυβ3 integrins) and CWR22 (αυβ3 integrin) cells to DU-145 and C4-2B cells that normally do not express integrin, which subsequently induces their progression and invasion [92,93]. More interestingly, exosomes are directly involved in the transport of chemotherapy resistance to other cells, as evidenced by a study conducted by Shedden et al. In this study, drug-resistant, cancer cell-associated exosomes were loaded with multidrug-resistant MDR-1 or p-gp proteins, which triggered the transfer of chemo-resistance to the recipient cells [76]. The exosomes released from the DU-145 and 22RV1 cells, which were resistant to doxorubicin, were not only able to transfer chemo-resistance to parental DU-145 and 22RV1 cells but also to LNCaP cells [94]. Furthermore, the impaired chemo-sensitivity to therapeutic agents is a reversible phenomenon in PCa. For example, when DU-145 cells with resistance to camptothecin or paclitaxel were treated with exosomes isolated from normal prostatic epithelial cells, including PrEC or the immortal cell line RWPE-1, their sensitivity was partially recovered [95,96]. Confirming this phenomenon, DU-145 cells with resistance to paclitaxel partially lost their resistance when treated with human mesenchymal stem cell (hMSC)-derived exosomes [96]. The different mechanisms by which exosomes opt to modify the genetic traits of PCa cells are outlined in Figure 1.
Regarding exosomal cargo, exosome-associated miRNAs are emerging as a novel regulator for many cellular functions, including cell metabolism. For example, miR-126, which is an angiogenesis inducer, regulates cancer metabolism via its downstream target insulin receptor substrate-1 (IRS1) [97,98]. IRS1 is not only a metabolic and growth-promoting protein via regulation of the insulin receptor (IR) and insulin-like growth factor I receptor (IGF-IR), but also contributes to neoplastic transformation [99]. Typically, endothelial cells (ECs) express and release miR-126 at a high level in the TME [100,101]. Notably, the level of exosome-associated miR-126 is elevated during glucose deprivation and oxidative stress [98,102,103]. In addition, the ectopic expression of miR-126 increases cellular glycolysis and mitochondrial dysfunction, downregulates Akt and FOXO1 signaling pathways, and upregulates gluconeogenesis and oxidative stress defense in malignant mesothelioma cells [97,98,104]. Cancer-associated exosomes containing miR-122 inhibit glycolytic metabolism by surrounding cells in the pre-metastatic niche via the regulation of pyruvate kinase muscle isozyme M2 (PKM2) and then allowing glucose uptake by growing cancer cells [105]. Further studies are warranted to validate the impact of PCa-derived exosomes in the metabolic crosstalk between cancer and stromal cells, which contributes not only to cell survival when oxygen and nutrients are deprived, but also to cancer progression and aggressiveness.

3.3. Effect of Exosomal Cargo on the Metabolic Reprogramming of Stromal Cells in the TME

Altered cell metabolism is a hallmark of cancer progression. According to the Warburg effect, cancer cells have a high tendency to ferment glucose, which is a process associated with lactate production, even in the presence of oxygen, that subsequently results in a reduced pH in the TME and, therefore, leads to cancer progression and aggressiveness [106]. There is increasing concern regarding the role of tumor-derived exosomes in the modulation of stromal cell metabolism. Inside the TME, metabolic coordination between cancer cells and stromal cells, such as cancer-associated fibroblasts (CAFs), tumor-associated macrophages (TAM), bone-marrow-derived cells (BMDCs), and tumor endothelial cells (TECs), plays a significant role in cancer cell survival and growth [107]. Most often, the adaptation of stromal cells to glycolysis supports cancer growth via the exchange of exosomes that provide the cancer cells with metabolic intermediates such as lactate, pyruvate, ketones, and glutamine, which can be used by cancer cells for the biosynthesis of macromolecules [107]. In this venue, fibroblasts, which represent one-third of stromal cells, are key players in cancer development [108,109,110]. Normal, dormant fibroblasts negatively affect the growth of tumor cells by maintaining epithelial homeostasis and proliferative latency [111,112]. Tumorigenic cells tend to induce pseudo-hypoxia by inducing HIFα, reactive oxygen species (ROS), and different oncogenic signals within the TME. These events are followed by the recruitment of normal-associated fibroblasts (NAFs) and their reprogramming into cancer-associated fibroblasts (CAFs), which are also known as myofibroblasts [113]. CAFs give-and-take both signaling molecules and metabolic fuels with the cancer cells, either by secreting individual molecules such as lactate or via the transfer of exosomal cargo. This process regulates the metabolic activities in the neighboring cancer cells and forcing them to respire and overcome their energy depletion [108,109,114]. Cancer-derived exosomes induce the Warburg effect and increase the rate of glycolysis, followed by lactate production in stromal cells [115]. One research team reported that CAF-derived exosomes promote a metabolic shift in PCa cells through the inhibition of mitochondrial oxidative phosphorylation. This caused an increase in glycolysis and the production of a variety of metabolites such as lactate, acetate, amino acids, tricarboxylic acid cycle intermediates, and lipids [116].

3.4. Desmoplastic Response of Stromal Cells to the PCa-Derived Exosomal Proteome

Essentially, the consequence of the mutual interaction between cancer cells and stromal cells on the TME favors cell survival, proliferation, angiogenesis, resistance to therapy, immune avoidance, and metastasis. Ample published data regarding the biology of exosomes have demonstrated that exosomes released by cancer cells reprogram their TME [117]. Undeniably, tumor-stromal cell crosstalk is always associated with tumor aggressiveness and the morphological transformation of stromal cells [118]. Proteomic studies of cancer-associated exosomes by advanced mass spectrometry have indicated that many cytoplasmic, membranous, Golgi apparatus, and endoplasmic reticulum proteins can be encapsulated in exosomal vesicles [119,120,121,122]. Cancer-associated exosomes opt to transport specific transmembrane proteins, including integrins and tetraspanins such as CD9, CD63, CD81, and CD82 that are specifically recognized by target cells, which can explain the high rate of exosomal uptake by cancer-adjacent stromal cells [117,123]. Interestingly, cancer-associated exosomes transport many functional proteins such as endosomal network proteins, which include (i) membrane transport and fusion proteins (GTPases, annexins, Rab proteins, and flotillin), (ii) heat-shock proteins (HSPs60, 70, and 90), (iii) multivesicular bodies (MVBs) biogenesis proteins (Alix and TSG101), (iv) cytoskeletal proteins (actin, tubulin, syntenin, and moesin), (v) lipoproteins and phospholipases, (vi) metabolic enzymes, and (vii) signal transduction proteins and major histocompatibility complement antigens [124,125,126,127].
In general, PCa-associated exosomes procured from clinical samples reveal cargo that contains cancer-related proteins such as CD9, CD81, and TSG101, Annexin A2, Fatty Acid Synthase (FASN), and prostate-specific membrane antigen (PSMA: a PCa-specific biomarker) [128,129]. PCa cell-derived exosomes transport Ras superfamily of GTPases Rab1a, Rab1b, and Rab11a to the TME, which then contribute to tumorigenic reprogramming and the recruitment of adipose-derived stem cells (ASCs), thereby supporting PCa cell growth and clonal expansion [130]. Emerging evidence shows that PCa-associated exosomes shuttle inactive TGFβ1, which induces a pro-tumorigenic phenotype differentiation of normal fibroblasts to CAFs via Mothers against Decapentaplegic Homolog 1 (SMAD)-dependent and -independent signaling pathways [131,132,133,134,135]. PCa-associated exosomes can also trigger the differentiation of bone marrow-mesenchymal stem cells (BM-MSC) to CAF cells, causing cells to become more active by producing high levels of VEGFA, HGF, and matrix metallopeptidase (MMP), which are associated with tumor growth [136]. Moreover, the treatment of cells with PCa-associated exosomal TGFβ1 increases the aggressive phenotype in CAFs compared to cells treated with the soluble form [132]. PCa-associated exosomes under hypoxic conditions contain nearly three times as much protein (CD63, CD81, HSP90, HSP70, Annexin II, TGF-β2, TNF1α, IL6, TSG101, Akt, ILK1, and β-catenin) as exosomes in normoxic conditions, which ultimately promotes CAF formation [91]. Furthermore, PCa-associated exosomes deliver the integrin αvβ3 to the TME where this integrin triggers the activation of Src phosphorylation and enhances the expression of pro-inflammatory S100 in stromal cells [93,137]. Src-family kinases are usually expressed in the prostatic epithelium, with their expression increasing during PCa initiation and progression, thus it is suggested that this kinase family is linked to normal cell transformation [138]. Many studies have demonstrated the vital role of c-Src tyrosine kinase, insulin-like growth factor 1 receptor (IGF-1R), and focal adhesion kinase (FAK) in PCa development and angiogenesis [139,140]. Remarkably, PCa-associated exosomes are enriched by c-Src, IGF-1R, and FAK proteins [141]. Aggressive PCa-associated exosomes contain urokinase-type plasminogen activator (uPA) [142], a stimulator for a protease plasminogen that is linked to vascular structure remodeling [143]. Moreover, PCa-associated exosomes promote the escape of tumors from immune surveillance by compromising the cytotoxic function of lymphocytes and reduce NKG2D receptor expression on natural killer and CD8+ T cells [144]. Cancer-associated exosomes not only shuttle immune regulatory molecules such as FasL, TGF-β, galectin-9, and HSP72, which help cancer cells escape the immune system, but also trigger the Fas/FasL pathway to induce CD8+ T cells toward the apoptotic pathway [145,146]. One recent study revealed that oncosomes (100–400 nm in diameter) in addition to exosomes can modify the prostate stroma phenotype [147]. Minciacchi et al. reported that the uptake of large oncosomes by prostate fibroblasts induces an αSMA-positive CAF phenotype that is independent of other CAFs markers such as MMP1, thrombospondin-1 (TSP-1), and TGFβ1, proving that oncosomes induce a distinct CAF phenotype [148], as outlined in Table 1 and Figure 2.

3.5. Exosomal Lipids as Messengers in PCA-Stromal Cell Crosstalk

Lipid metabolism is overwhelmingly disturbed in cancer cells compared to normal cells. Exosomes reflect the composition of different lipid types in cancer cells. Lipids have been implicated in different aspects of exosome biogenesis and various functions. The exosomal membrane is a lipid bilayer enriched in lipid raft-like domains. These domains act as platforms for lipid raft-associated proteins [161], diglycerides, sphingolipids, and glycerophospholipids including phosphatidylcholine (PC), phosphatidylserine (PS), phosphatidylethanolamines (PE), and phosphatidylinositol (PI) [162]. Of note, exosomes gain their rigidity from their lipid composition, which gives them stability in extracellular fluids, and also facilitates the process of internalization by recipient cells [162,163]. Although similarities exist in the orientations of membranous molecules between the exosomal membrane and their parental cell membranes, nonetheless the distribution of PS in the exosomal membrane (enriched in the outer leaflet) is in contrast to the distribution of these molecules in the membrane of live parental cells (inner leaflet) [162]. It has been shown that exosomal prostaglandins (PGE2) are involved in immunosuppression and can trigger tumor growth [164]. Interestingly, the transfer of phosphatidylcholine transporter ABCA3 by exosomes has been indicated in B lymphoma treatment that is resistant to immunotherapy with rituximab [79]. Moreover, exosomal lipids induce apoptosis in SOJ-6 human pancreatic cancer cells via inhibition of the Notch-1 pathway [165]. Exosomal lipids also increase drug resistance in human pancreatic cancer MiaPaCa-2 cells through the C-X-C motif chemokine receptor 4 (CXCR4)/stromal cell derived factor (SDF)-1α signaling pathway [166]. Finally, exosomal lipids can be considered as biomarkers in PCa [167]. However, further studies are warranted to define the lipid composition of PCa-associated exosomes and whether the lipid composition becomes altered at different tumor stages, which could affect the biological functions of exosomes in stromal cells

3.6. The Role of Exosomes in Shuttling Bioactive Materials between PCA and Stromal Cells

Exosome-mediated plane transfer of genetic and epigenetic materials comprising multiple RNA species such as messenger RNA (mRNA), microRNAs (miRNAs), long noncoding RNAs (lncRNAs), ribosomal RNA (rRNA), piwi RNA (piRNAs), small-nuclear RNA (snRNA), small-nucleolar RNA (snoRNA), and transfer RNA as well as genomic and mitochondrial DNA among cancer cells and the stromal cells are significant pathways for maintaining a favorable environment for cancer growth [66,168,169]. miRNAs are evolutionarily known as short, noncoding, single-stranded RNAs of 22 nucleotides in length with partial homology to sequences in their target mRNA. miRNAs regulate the posttranscriptional level of gene expression via the formation of a silencing complex (RISC), which allows annealing of miRNAs to the 3′UTR target genes and consequently represses protein expression. The second choice of posttranscriptional regulation occurs through mRNA destabilization [170,171,172]. Emerging evidence has shown that exosomal miRNAs regulate many biological functions in the TME such as the desmoplastic response of stromal cells and the proliferation, apoptosis, and invasion of tumor cells [173]. One study revealed that PCa cell-associated exosomes deliver onco-miRNAs including miR-125b, miR-130b, and miR-155 and onco-mRNAs such as H-ras and K-RAS transcripts, which have a desmoplastic effect on ASCs, thereby supporting tumor progression and colonization [130]. Moreover, miR-155 can suppress the expression of its target protein 53-induced nuclear protein 1 (TP53INP1) to promote CAF-like phenotypes in fibroblasts [174].
The cargo of PCa cell-derived exosomes not only induces the expression of prostate-specific genes in bone marrow cells (BMCs) but also in normal human cells [175]. More specifically, PCa-derived exosomes transport miR-100, miR-21, and miR-139, which upregulate the expression of receptor activator of nuclear factor kappa-Β ligand (RANKL) and Metalloproteinases in CAFs and promote PCa growth and metastasis [176]. Metastasis-initiating cells (MICs) are distinct circulating tumor cells (CTCs) with a strong ability to grow, survive, and colonize PCa cells in distant metastatic organs [177]. MIC-derived exosomes can modify the TME to stimulate the oncogenic transformation of normal epithelial and stromal cells, which stimulate phenotype transformation and promote PCa epithelial-mesenchymal transition (EMT) via the activation of RANKL, FOXM1, and c-Myc [177,178,179]. In addition, metastatic PCa patient-derived exosomes express a high level of miR-141 and miR-21, which regulate osteoclastogenesis and osteoblastogenesis [180,181]. One group of researchers showed that PC-3-derived exosomes regulate osteoclastogenesis and osteoblast proliferation and promote bone metastasis [182]. Another research group revealed that EVs transfer miR-409, miR-379, and miR-154, which have a vital role in embryogenesis and pluripotent stem cell formation, and favor PCa carcinogenesis and metastasis by activating tumor-stroma interactions [177].
Long non-coding RNAs (lncRNA), measuring more than 200 nucleotides in length, have a potential role in many physiological and pathophysiological processes [183]. In a comparative study using the lncRNA array, PCa cell line-associated exosomes overexpress 26 lncRNAs compared to normal epithelial cells [184]. These exosomal lncRNAs are significantly enriched in target motifs for the miRs, which are found in the same exosomes, suggesting that the sorting of exosomes is a highly organized process that allows for the selection of specific miRs and lncRNAs to be uploaded in their vesicles [184]. It has been reported that exosome-associated lncRNA MYU induces adjacent PCa cell proliferation and migration by competitively binding to miR-184 and therefore upregulates c-Myc [185]. Moreover, urine-derived exosomes collected from PCa patients are enriched in lncRNA-p21, which is associated with the malignancy of PCa [186]. In addition to the reported studies summarized in Table 2 and Figure 2, the role of PCa-derived exosomal lncRNA cargo in the recruitment of stromal cells that are still in the growth phase remains to be understood, such that further studies are needed to fully address this phenomenon.

3.7. Two-Way Crosstalk between Stromal Cells and PCA Cell-Associated Exosomal Cargo

It is noteworthy, that PCa-associated CAFs promote PCa progression, metastasis and development of resistance to therapy [191,192], as summarized in Figure 3.
Several mechanisms are suggested to explain the possible role of CAFs in promoting PCa progression. The first mechanism anticipates the role of CAFs in supporting PCa progression by increasing extracellular matrix (ECM) deposition and turnover in CAFs, which are accompanied by the production of cytokines such as TGF-β [193]. The second mechanism arises due to the overexpression of CAF growth and angiogenic factors, such as growth/differentiation factor 15 (GDF15) [194]. The third mechanism is a significant increase in the release of soluble factors and insoluble ECMs by CAFs, which promote the neoplastic transformation of normal cells to PCa-like cells [195]. In addition, CAF-derived exosomes transfer miRNAs into neighboring epithelial cells, which caused tumor growth in a PCa-mouse model through the EMT pathway [196,197]. CAF-derived exosomes transfer miRNAs and proteins, including miR-21, miR-409, and CD81 to adjacent epithelial cells and promote cell proliferation, invasion, and chemo-resistance and alter metabolic pathways in PCa cells [145,196,197]. Interestingly, miR-21 represses the expression of its target’s apoptotic peptidase activating factor 1 (APAF1) and programmed cell death 4 (PDCD4) to inhibit apoptosis and increase paclitaxel resistance in cancer cells [198]. In the same context, osteoblast-derived exosomes can promote PC-3 cell proliferation [182]. In connection to the previous studies, the subcutaneous co-injection of ARCaPE cells (which naturally lack miR-409) in combination with miR-409-expressing stromal fibroblast cells promoted ARCaPE cell proliferation when compared to control cells, suggesting that exosomal miR-409 can be transferred from stromal cells to PCa cells [196]. A similar study using exosomes isolated from menstrual stem cells (MenSCs) to treat PC-3 cells induced the inhibition of VEGF on mRNA and protein levels, decreased ROS production, and the secretion of HIFα and, therefore, suppressed PCa cell growth [199].

4. The Immune-Modulatory Role of Exosomes

The immune system is one of the main barriers to develop cancer cells. Hence, complex communication between immune cells and growing tumor cells is crucial for cancer initiation, progression, and metastasis. Delineating the role of exosomes in regulating immune cells during and after cancer development is needed to develop advanced exosome-based vaccines and immune therapy. Neoplastic modulations of the immune cells include up-regulation of precise genes and proteins and subsequent escape from immune cell recognition and killing [200,201,202]. Inside TME, cancer cells are able to remodel cytotoxic T lymphocytes (CTLs) and natural killer (NK) cells to facilitate tumor progression [203,204]. It was reported that cancer-associated exosomes interfere with the development of CD14+ monocytes into mature dendritic cells (DCs) [205]. More interestingly, the activated myeloid-derived suppressor cells (MDSCs) decreased NK and CD4+/CD8+ lymphocytes and reduced their cytotoxic effect through the interaction between exosomal HSP72, TLR-2, and MyD88 and immune cells [81]. Cancer-associated exosomes not only shuttle immune regulatory molecules such as PD-L1, TRAIL, TGF-β, IL-10, FasL, galectin-9, HSP72, and PGE2 to exert an immunosuppressive effect, but they also to trigger the Fas/FasL pathway and induce apoptosis in CD8+ T cells [145,146,206,207,208,209,210]. Cancer-associated exosomes can also alter the behavior of macrophages, which regulate host immunity. This promotes tumor progression by releasing cytokines, inducing tissue remodeling, and promoting angiogenesis and metastasis [211,212,213]. Moreover, PCa-associated exosomes promote tumor escape from immune surveillance by compromising the cytotoxic function of lymphocytes and reducing NKG2D receptor expression in NK and CD8+ T cells [144]. On the contrary, circulating NK and CD8+ T cells in CRPC patients decrease surface killer cell lectin-like receptor K1 (KLRK1) expression. Therefore, treating healthy lymphocytes with CRPC serum-derived exosomes decreases the expression of KLRK1 in effector lymphocytes [144]. In the same scenario, cancer-associated exosomes shuttle particular miRNAs such as miR-24-3p, miR-891a, miR-106a-5p, miR-20a-5p, and miR-1908, which compromise T-cell function in nasopharyngeal cancer [214,215]. Furthermore, pancreatic cancer-associated exosomes shuttle miR-212-3p to inhibit regulatory factor X-associated protein (RFXAP), a regulatory transcription factor for the MHC-II gene, and initiate immune tolerance in dendritic cells (DCs) [216]. In addition, an in vivo study conducted on pancreatic cancer revealed that exosomal miR-203 downregulates the expression of TRL4, affects TNF-α and IL-1 production, and deteriorates DC development and Th1 differentiation [217]. In another study, engineered exosomes with TNF-related apoptosis-inducing ligand (TRAIL) rewired the apoptotic pathway in vitro and in a preclinical mouse model [218].

5. Targeting the Tumor Microenvironment

TMEs share common features such as hypoxia, oxidative stress and acidosis, uncontrolled growth, a resistance to apoptosis, a metabolic shift toward anaerobic glycolysis and the remodeling of ECM-associated cells [219]. In addition to other biological factors, exosomes secreted by cancer and TME cells modulate stromal cells residing in the TME [220,221]. Understanding such changes in the TME during tumor progression will yield additional advantages in the development of novel therapeutic strategies to tackle cancer progression and metastasis compared to the currently available options, for example, by targeting the ECM by using angiotensin II, TGF-β, and heparanase inhibitor roneparstat (SST0001) [222,223,224,225,226], affecting hypoxia and acidosis by targeting hypoxia-induced factor-1 (HIF-1) using several compounds and therapies such as Topotecan [227,228,229], and targeting endothelial cells and pericytes to avoid neovascularization using several antiangiogenic drugs such as bevacizumab (Avastin) [230,231]. In addition, targeting the recruitment of tumor-associated-macrophages (TMAs) in the TME [232], activating the anti-tumor activity of immune cells [233], and targeting CAFs [230] might add new therapeutic benefits in the treatment of tumors in advanced stages. In the same context, targeting cancer-associated exosomes may open new venues in the treatment of aggressive forms of PCa. Toward this direction, Bastos and colleagues discussed different means of targeting exosomes during the process of exosomal biogenesis, release, and uptake [234]. Tumor-associated exosomes can be targeted during the process of biogenesis and released by inhibiting ceramides using the GW4869 inhibitor [235] and Rab-associated proteins [236]. Manumycin A blocks biogenesis and secretion of exosomes by inhibiting the activity of ERK1/2 and expression of heterogeneous nuclear ribonucleoprotein H1 (hnRNPH1) in CRPC cells [237]. In addition, blocking the mechanisms of exosome internalization is another approach for targeting cancer-associated cells in the TME. For example, methyl-β-cyclodextrin (MCβD) inhibits exosome uptake in a number of cancer cells by disrupting the lipid rafts of exosomes [238]. Cytochalasin D, heparin, dynasore, and nystatin are other examples of exosome uptake inhibitors [234]. However, mechanisms of targeting exosomes are very complicated because some of these pathways are involved in most cellular activities. Therefore, further studies are warranted to shed light on the biology of exosomes to find more specific targets.

6. Conclusions and Future Directions

The advancement of science and technology opens new venues for understanding new mechanisms underlying cancer initiation, progression, and metastasis and, therefore, offers new treatment options for improving the survival of cancer patients. One important scientific direction in recent years has been to shed light on the various roles of exosomes in cancer biology, in addition to their clinical applications. The mechanisms by which tumor cells send and receive molecular signals from the TME is a growing area of research. A significant quantity of research has indicated that exosomal cargo is a powerful communication means in cell-cell interactions, and is an integral part of the crosstalk between tumor and stromal cells in the TME. Given that exosomes contribute to cancer progression through the transfer of different types of exosomal cargo to their target cells, these bioactive molecules stimulate multiple oncogenic pathways, which remodel normal, cancer, and stromal cells inside the TME. Elucidating the underlying mechanisms of cancer aggressiveness by shuttling exosomal cargo between PCa and stromal cells in the TME increases the viability of using exosomes as promising therapeutic agents. In addition, these vesicles can serve as vectors for many anticancer agents, a) because of their biological nature and cellular origin, b) because of lipophilic nature that makes them able to easily cross membranes such as the blood-brain barrier, c) because they are not filtered by the glomerulus, which gives exosomes long circulation lifetimes, and d) because exosomes can carry different types of bioactive molecules. Exosomal cargo can be considered as potential diagnostic and prognostic markers for different types of cancers, which has created excitement in the scientific community because this fact paves the way for using exosomal cargo as reliable tumor markers. Another application is to use exosomes to unwire the connection between cancer cells and the TMEs, with the potential to reduce PCa metastasis and the development of drug resistance. Precise information regarding how exosomes selectively load their cargo, independent of their mother cells, and then release and internalize these bioactive molecules into target cells will reshape our treatment modalities for cancer patients. Although a growing number of institutes and agencies share a wealth of biological data on public domains, more collaborative studies and access to these exosomal data remain in demand. The assembly of multidisciplinary research teams is highly encouraged to understand the biology of the TME and clinical utilities of exosomes in the context of cancer disease and its effective treatment.

Author Contributions

Conceptualization, Z.Y.A.E., H.I.A. and M.Z.; methodology, S.H.S., H.E.A.A., R.G., and M.G.; software, S.H.S.; validation, S.H.S., H.I.A., M.Z. and Z.Y.A.E.; investigation, S.H.S., Z.Y.A.E.; resources, S.H.S., Z.Y.A.E.; data curation, S.H.S., H.E.A.A., Z.Y.A.E.; writing—original draft preparation, S.H.A., H.E.A.A., R.G., M.G.; writing—review and editing, S.H.A., H.E.A.A., H.I.A., M.Z., Z.Y.A.E.; visualization, S.H.A., Z.Y.A.E.; supervision, Z.Y.A.E.; project administration, Z.Y.A.E.; funding acquisition, Z.Y.A.E. All authors have read and agreed to the published version of the manuscript.


This study was partially supported by the NIH grant number R21CA194750 (ZYA), and the Texas A&M Health Science Center (ZYA).

Conflicts of Interest

The authors declare there are no conflict of interest.


  1. Minciacchi, V.; Zijlstra, A.; Rubin, M.A.; Di Vizio, D. Extracellular vesicles for liquid biopsy in prostate cancer: Where are we and where are we headed? Prostate Cancer Prostatic Dis. 2017, 20, 251. [Google Scholar] [CrossRef] [PubMed]
  2. Shukla, M.E.; Yu, C.; Reddy, C.A.; Stephans, K.L.; Klein, E.A.; Abdel-Wahab, M.; Ciezki, J.; Tendulkar, R.D. Evaluation of the current prostate cancer staging system based on cancer-specific mortality in the surveillance epidemiology and end results database. Clin. Genitourin Cancer 2015, 13, 17–21. [Google Scholar] [CrossRef] [PubMed]
  3. American Cancer Society. Cancer Facts & Figures 2019; American Cancer Society: Atlantaga, GA, USA, 2019. [Google Scholar]
  4. Endzeliņš, E.; Melne, V.; Kalniņa, Z.; Lietuvietis, V.; Riekstiņa, U.; Llorente, A.; Linē, A. Diagnostic, prognostic and predictive value of cell-free miRNAs in prostate cancer: A systematic review. Mol. Cancer 2016, 15, 41. [Google Scholar] [CrossRef] [PubMed]
  5. Hutchinson, L. Closing the controversies gap in prostate cancer? Nat. Rev. Clin. Oncol. 2014, 11, 299. [Google Scholar] [CrossRef]
  6. Cheville, J.C.; Tindall, D.; Boelter, C.; Jenkins, R.; Lohse, C.M.; Pankratz, V.S.; Sebo, T.J.; Davis, B.; Blute, M.L. Metastatic prostate carcinoma to bone: Clinical and pathologic features associated with cancer-specific survival. Cancer 2002, 95, 1028–1036. [Google Scholar] [CrossRef]
  7. Scher, H.I.; Halabi, S.; Tannock, I.; Morris, M.; Sternberg, C.N.; Carducci, M.A.; Eisenberger, M.A.; Higano, C.; Bubley, G.J.; Dreicer, R. Design and end points of clinical trials for patients with progressive prostate cancer and castrate levels of testosterone: Recommendations of the Prostate Cancer Clinical Trials Working Group. J. Clin. Oncol. Off. J. Am. Soc. Clin. Oncol. 2008, 26, 1148. [Google Scholar] [CrossRef]
  8. Chen, C.D.; Welsbie, D.S.; Tran, C.; Baek, S.H.; Chen, R.; Vessella, R.; Rosenfeld, M.G.; Sawyers, C.L. Molecular determinants of resistance to antiandrogen therapy. Nat. Med. 2004, 10, 33. [Google Scholar] [CrossRef]
  9. Debes, J.D.; Tindall, D.J. Mechanisms of androgen-refractory prostate cancer. New Engl. J. Med. 2004, 351, 1488–1490. [Google Scholar] [CrossRef]
  10. Hu, R.; Dunn, T.A.; Wei, S.; Isharwal, S.; Veltri, R.W.; Humphreys, E.; Han, M.; Partin, A.W.; Vessella, R.L.; Isaacs, W.B. Ligand-independent androgen receptor variants derived from splicing of cryptic exons signify hormone-refractory prostate cancer. Cancer Res. 2009, 69, 16–22. [Google Scholar] [CrossRef]
  11. Stanbrough, M.; Bubley, G.J.; Ross, K.; Golub, T.R.; Rubin, M.A.; Penning, T.M.; Febbo, P.G.; Balk, S.P. Increased expression of genes converting adrenal androgens to testosterone in androgen-independent prostate cancer. Cancer Res. 2006, 66, 2815–2825. [Google Scholar] [CrossRef]
  12. Holzbeierlein, J.; Lal, P.; LaTulippe, E.; Smith, A.; Satagopan, J.; Zhang, L.; Ryan, C.; Smith, S.; Scher, H.; Scardino, P. Gene expression analysis of human prostate carcinoma during hormonal therapy identifies androgen-responsive genes and mechanisms of therapy resistance. Am. J. Pathol. 2004, 164, 217–227. [Google Scholar] [CrossRef]
  13. Agarwal, N.; Hutson, T.E.; Vogelzang, N.J.; Sonpavde, G. Abiraterone acetate: A promising drug for the treatment of castration-resistant prostate cancer. Future Oncol. 2010, 6, 665–679. [Google Scholar] [CrossRef] [PubMed]
  14. Sonpavde, G.; Attard, G.; Bellmunt, J.; Mason, M.D.; Malavaud, B.; Tombal, B.; Sternberg, C.N. The role of abiraterone acetate in the management of prostate cancer: A critical analysis of the literature. Eur. Urol. 2011, 60, 270–278. [Google Scholar] [CrossRef] [PubMed]
  15. Attard, G.; Reid, A.H.; Auchus, R.J.; Hughes, B.A.; Cassidy, A.M.; Thompson, E.; Oommen, N.B.; Folkerd, E.; Dowsett, M.; Arlt, W. Clinical and biochemical consequences of CYP17A1 inhibition with abiraterone given with and without exogenous glucocorticoids in castrate men with advanced prostate cancer. J. Clin. Endocrinol. Metab. 2012, 97, 507–516. [Google Scholar] [CrossRef] [PubMed]
  16. Attard, G.; Reid, A.H.; A’Hern, R.; Parker, C.; Oommen, N.B.; Folkerd, E.; Messiou, C.; Molife, L.R.; Maier, G.; Thompson, E. Selective inhibition of CYP17 with abiraterone acetate is highly active in the treatment of castration-resistant prostate cancer. J. Clin. Oncol. Off. J. Am. Soc. Clin. Oncol. 2009, 27, 3742. [Google Scholar] [CrossRef] [PubMed]
  17. Matsunaga, N.; Kaku, T.; Ojida, A.; Tanaka, T.; Hara, T.; Yamaoka, M.; Kusaka, M.; Tasaka, A. C17, 20-lyase inhibitors. Part 2: Design, synthesis and structure–activity relationships of (2-naphthylmethyl)-1H-imidazoles as novel C17, 20-lyase inhibitors. Bioorganic Med. Chem. 2004, 12, 4313–4336. [Google Scholar] [CrossRef]
  18. Niraula, S.; Chi, K.; Joshua, A.M. Beyond Castration—Defining Future Directions in the Hormonal Treatment of Prostate Cancer. Horm. Cancer 2012, 3, 3–13. [Google Scholar] [CrossRef]
  19. Taplin, M.-E.; Montgomery, R.B.; Group, A. ARMOR2: Galeterone in progressive CRPC patients who have failed oral therapy. Am. Soc. Clin. Oncol. 2014, 32, 71. [Google Scholar] [CrossRef]
  20. Tran, C.; Ouk, S.; Clegg, N.J.; Chen, Y.; Watson, P.A.; Arora, V.; Wongvipat, J.; Smith-Jones, P.M.; Yoo, D.; Kwon, A. Development of a second-generation antiandrogen for treatment of advanced prostate cancer. Science 2009, 324, 787–790. [Google Scholar] [CrossRef]
  21. Clegg, N.J.; Wongvipat, J.; Joseph, J.D.; Tran, C.; Ouk, S.; Dilhas, A.; Chen, Y.; Grillot, K.; Bischoff, E.D.; Cai, L. ARN-509: A novel antiandrogen for prostate cancer treatment. Cancer Res. 2012, 72, 1494–1503. [Google Scholar] [CrossRef]
  22. Kirby, M.; Hirst, C.; Crawford, E. Characterising the castration-resistant prostate cancer population: A systematic review. Int. J. Clin. Pract. 2011, 65, 1180–1192. [Google Scholar] [CrossRef] [PubMed]
  23. Giannakakou, P.; Nakano, M.; Nicolaou, K.C.; O’Brate, A.; Yu, J.; Blagosklonny, M.V.; Greber, U.F.; Fojo, T. Enhanced microtubule-dependent trafficking and p53 nuclear accumulation by suppression of microtubule dynamics. Proc. Natl. Acad. Sci. USA 2002, 99, 10855–10860. [Google Scholar] [CrossRef] [PubMed]
  24. Tannock, I.F.; De Wit, R.; Berry, W.R.; Horti, J.; Pluzanska, A.; Chi, K.N.; Oudard, S.; Théodore, C.; James, N.D.; Turesson, I. Docetaxel plus prednisone or mitoxantrone plus prednisone for advanced prostate cancer. New Engl. J. Med. 2004, 351, 1502–1512. [Google Scholar] [CrossRef] [PubMed]
  25. Mita, A.C.; Denis, L.J.; Rowinsky, E.K.; DeBono, J.S.; Goetz, A.D.; Ochoa, L.; Forouzesh, B.; Beeram, M.; Patnaik, A.; Molpus, K. Phase I and pharmacokinetic study of XRP6258 (RPR 116258A), a novel taxane, administered as a 1-h infusion every 3 weeks in patients with advanced solid tumors. Clin. Cancer Res. 2009, 15, 723–730. [Google Scholar] [CrossRef] [PubMed]
  26. Taylor, B.; Varambally, S.; Chinnaiyan, A. Differential proteomic alterations between localised and metastatic prostate cancer. Br. J. Cancer 2006, 95, 425. [Google Scholar] [CrossRef]
  27. Joniau, S.; Abrahamsson, P.-A.; Bellmunt, J.; Figdor, C.; Hamdy, F.; Verhagen, P.; Vogelzang, N.J.; Wirth, M.; Van Poppel, H.; Osanto, S. Current vaccination strategies for prostate cancer. Eur. Urol. 2012, 61, 290–306. [Google Scholar] [CrossRef]
  28. Bissell, M.J.; Hines, W.C. Why don’t we get more cancer? A proposed role of the microenvironment in restraining cancer progression. Nat. Med. 2011, 17, 320. [Google Scholar] [CrossRef]
  29. Wiseman, B.S.; Werb, Z. Stromal effects on mammary gland development and breast cancer. Science 2002, 296, 1046–1049. [Google Scholar] [CrossRef]
  30. Egeblad, M.; Nakasone, E.S.; Werb, Z. Tumors as organs: Complex tissues that interface with the entire organism. Dev. Cell 2010, 18, 884–901. [Google Scholar] [CrossRef]
  31. Shephard, A.P.; Yeung, V.; Clayton, A.; Webber, J.P. Prostate cancer exosomes as modulators of the tumor microenvironment. J. Cancer Metastasis Treat. 2017, 3, 288–301. [Google Scholar] [CrossRef]
  32. Saleem, S.N.; Abdel-Mageed, A.B. Tumor-derived exosomes in oncogenic reprogramming and cancer progression. Cell Mol. Life Sci 2015, 72, 1–10. [Google Scholar] [CrossRef] [PubMed]
  33. Akers, J.C.; Gonda, D.; Kim, R.; Carter, B.S.; Chen, C.C. Biogenesis of extracellular vesicles (EV): Exosomes, microvesicles, retrovirus-like vesicles, and apoptotic bodies. J. Neuro-Oncol. 2013, 113, 1–11. [Google Scholar] [CrossRef] [PubMed]
  34. Riches, A.; Campbell, E.; Borger, E.; Powis, S. Regulation of exosome release from mammary epithelial and breast cancer cells–a new regulatory pathway. Eur. J. Cancer 2014, 50, 1025–1034. [Google Scholar] [CrossRef] [PubMed]
  35. Wei, Y.; Wang, D.; Jin, F.; Bian, Z.; Li, L.; Liang, H.; Li, M.; Shi, L.; Pan, C.; Zhu, D. Pyruvate kinase type M2 promotes tumour cell exosome release via phosphorylating synaptosome-associated protein 23. Nat. Commun. 2017, 8, 14041. [Google Scholar] [CrossRef]
  36. Liu, C.; Su, C. Design strategies and application progress of therapeutic exosomes. Theranostics 2019, 9, 1015–1028. [Google Scholar] [CrossRef]
  37. Mashouri, L.; Yousefi, H.; Aref, A.R.; Ahadi, A.M.; Molaei, F.; Alahari, S.K. Exosomes: Composition, biogenesis, and mechanisms in cancer metastasis and drug resistance. Mol. Cancer 2019, 18, 75. [Google Scholar] [CrossRef]
  38. Robbins, P.D.; Morelli, A.E. Regulation of immune responses by extracellular vesicles. Nat. Rev. Immunol. 2014, 14, 195. [Google Scholar] [CrossRef]
  39. Koliopanos, A.; Friess, H.; Kleeff, J.; Shi, X.; Liao, Q.; Pecker, I.; Vlodavsky, I.; Zimmermann, A.; Büchler, M.W. Heparanase expression in primary and metastatic pancreatic cancer. Cancer Res. 2001, 61, 4655–4659. [Google Scholar]
  40. Koo, T.H.; Lee, J.-J.; Kim, E.-M.; Kim, K.-W.; Do Kim, H.; Lee, J.-H. Syntenin is overexpressed and promotes cell migration in metastatic human breast and gastric cancer cell lines. Oncogene 2002, 21, 4080. [Google Scholar] [CrossRef]
  41. Liu, R.-T.; Huang, C.-C.; You, H.-L.; Chou, F.-F.; Hu, C.-C.A.; Chao, F.-P.; Chen, C.-M.; Cheng, J.-T. Overexpression of tumor susceptibility gene TSG101 in human papillary thyroid carcinomas. Oncogene 2002, 21, 4830. [Google Scholar] [CrossRef]
  42. Oh, K.; Stanton, M.; West, W.; Todd, G.; Wagner, K. Tsg101 is upregulated in a subset of invasive human breast cancers and its targeted overexpression in transgenic mice reveals weak oncogenic properties for mammary cancer initiation. Oncogene 2007, 26, 5950. [Google Scholar] [CrossRef] [PubMed]
  43. Toyoshima, M.; Tanaka, N.; Aoki, J.; Tanaka, Y.; Murata, K.; Kyuuma, M.; Kobayashi, H.; Ishii, N.; Yaegashi, N.; Sugamura, K. Inhibition of tumor growth and metastasis by depletion of vesicular sorting protein Hrs: Its regulatory role on E-cadherin and β-catenin. Cancer Res. 2007, 67, 5162–5171. [Google Scholar] [CrossRef] [PubMed]
  44. Morgan-Fisher, M.; Wewer, U.M.; Yoneda, A. Regulation of ROCK activity in cancer. J. Histochem. Cytochem. 2013, 61, 185–198. [Google Scholar] [CrossRef]
  45. Ruiz-Martinez, M.; Navarro, A.; Marrades, R.M.; Viñolas, N.; Santasusagna, S.; Muñoz, C.; Ramírez, J.; Molins, L.; Monzo, M. YKT6 expression, exosome release, and survival in non-small cell lung cancer. Oncotarget 2016, 7, 51515. [Google Scholar] [CrossRef] [PubMed]
  46. Al-Nedawi, K.; Meehan, B.; Micallef, J.; Lhotak, V.; May, L.; Guha, A.; Rak, J. Intercellular transfer of the oncogenic receptor EGFRvIII by microvesicles derived from tumour cells. Nat. Cell Biol. 2008, 10, 619. [Google Scholar] [CrossRef] [PubMed]
  47. Takasugi, M.; Okada, R.; Takahashi, A.; Chen, D.V.; Watanabe, S.; Hara, E. Small extracellular vesicles secreted from senescent cells promote cancer cell proliferation through EphA2. Nat. Commun. 2017, 8, 15729. [Google Scholar] [CrossRef] [PubMed]
  48. Imjeti, N.S.; Menck, K.; Egea-Jimenez, A.L.; Lecointre, C.; Lembo, F.; Bouguenina, H.; Badache, A.; Ghossoub, R.; David, G.; Roche, S. Syntenin mediates SRC function in exosomal cell-to-cell communication. Proc. Natl. Acad. Sci. USA 2017, 114, 12495–12500. [Google Scholar] [CrossRef]
  49. Parolini, I.; Federici, C.; Raggi, C.; Lugini, L.; Palleschi, S.; De Milito, A.; Coscia, C.; Iessi, E.; Logozzi, M.; Molinari, A. Microenvironmental pH is a key factor for exosome traffic in tumor cells. J. Biol. Chem. 2009, 284, 34211–34222. [Google Scholar] [CrossRef]
  50. King, H.W.; Michael, M.Z.; Gleadle, J.M. Hypoxic enhancement of exosome release by breast cancer cells. BMC Cancer 2012, 12, 421. [Google Scholar] [CrossRef]
  51. Li, L.; Li, C.; Wang, S.; Wang, Z.; Jiang, J.; Wang, W.; Li, X.; Chen, J.; Liu, K.; Li, C. Exosomes derived from hypoxic oral squamous cell carcinoma cells deliver miR-21 to normoxic cells to elicit a prometastatic phenotype. Cancer Res. 2016, 76, 1770–1780. [Google Scholar] [CrossRef]
  52. Wang, T.; Gilkes, D.M.; Takano, N.; Xiang, L.; Luo, W.; Bishop, C.J.; Chaturvedi, P.; Green, J.J.; Semenza, G.L. Hypoxia-inducible factors and RAB22A mediate formation of microvesicles that stimulate breast cancer invasion and metastasis. Proc. Natl. Acad. Sci. USA 2014, 111, E3234–E3242. [Google Scholar] [CrossRef] [PubMed]
  53. Yu, X.; Harris, S.L.; Levine, A.J. The regulation of exosome secretion: A novel function of the p53 protein. Cancer Res. 2006, 66, 4795–4801. [Google Scholar] [CrossRef] [PubMed]
  54. Kosaka, N.; Iguchi, H.; Yoshioka, Y.; Takeshita, F.; Matsuki, Y.; Ochiya, T. Secretory mechanisms and intercellular transfer of microRNAs in living cells. J. Biol. Chem. 2010, 285, 17442–17452. [Google Scholar] [CrossRef] [PubMed]
  55. Mittelbrunn, M.; Gutiérrez-Vázquez, C.; Villarroya-Beltri, C.; González, S.; Sánchez-Cabo, F.; González, M.Á.; Bernad, A.; Sánchez-Madrid, F. Unidirectional transfer of microRNA-loaded exosomes from T cells to antigen-presenting cells. Nat. Commun. 2011, 2, 282. [Google Scholar] [CrossRef]
  56. Trajkovic, K.; Hsu, C.; Chiantia, S.; Rajendran, L.; Wenzel, D.; Wieland, F.; Schwille, P.; Brugger, B.; Simons, M. Ceramide triggers budding of exosome vesicles into multivesicular endosomes. Science 2008, 319, 1244–1247. [Google Scholar] [CrossRef]
  57. Villarroya-Beltri, C.; Baixauli, F.; Gutiérrez-Vázquez, C.; Sánchez-Madrid, F.; Mittelbrunn, M. Sorting it out: Regulation of exosome loading. Semin. Cancer Biol. 2014, 28, 3–13. [Google Scholar] [CrossRef]
  58. Shen, B.; Wu, N.; Yang, J.-M.; Gould, S.J. Protein targeting to exosomes/microvesicles by plasma membrane anchors. J. Biol. Chem. 2011, 286, 14383–14395. [Google Scholar] [CrossRef]
  59. Villarroya-Beltri, C.; Gutiérrez-Vázquez, C.; Sánchez-Cabo, F.; Pérez-Hernández, D.; Vázquez, J.; Martin-Cofreces, N.; Martinez-Herrera, D.J.; Pascual-Montano, A.; Mittelbrunn, M.; Sánchez-Madrid, F. Sumoylated hnRNPA2B1 controls the sorting of miRNAs into exosomes through binding to specific motifs. Nat. Commun. 2013, 4, 2980. [Google Scholar] [CrossRef]
  60. Guduric-Fuchs, J.; O’Connor, A.; Camp, B.; O’Neill, C.L.; Medina, R.J.; Simpson, D.A. Selective extracellular vesicle-mediated export of an overlapping set of microRNAs from multiple cell types. Bmc Genom. 2012, 13, 357. [Google Scholar] [CrossRef]
  61. Squadrito, M.L.; Baer, C.; Burdet, F.; Maderna, C.; Gilfillan, G.D.; Lyle, R.; Ibberson, M.; De Palma, M. Endogenous RNAs modulate microRNA sorting to exosomes and transfer to acceptor cells. Cell Rep. 2014, 8, 1432–1446. [Google Scholar] [CrossRef]
  62. Rajagopal, C.; Harikumar, K. The origin and functions of exosomes in cancer. Front. Oncol. 2018, 8, 66. [Google Scholar] [CrossRef] [PubMed]
  63. Jansen, F.H.; Krijgsveld, J.; van Rijswijk, A.; van den Bemd, G.-J.; van den Berg, M.S.; van Weerden, W.M.; Willemsen, R.; Dekker, L.J.; Luider, T.M.; Jenster, G. Exosomal secretion of cytoplasmic prostate cancer xenograft-derived proteins. Mol. Cell. Proteom. 2009, 8, 1192–1205. [Google Scholar] [CrossRef]
  64. Nilsson, J.; Skog, J.; Nordstrand, A.; Baranov, V.; Mincheva-Nilsson, L.; Breakefield, X.; Widmark, A. Prostate cancer-derived urine exosomes: A novel approach to biomarkers for prostate cancer. Br. J. Cancer 2009, 100, 1603. [Google Scholar] [CrossRef] [PubMed]
  65. Duijvesz, D.; Luider, T.; Bangma, C.H.; Jenster, G. Exosomes as biomarker treasure chests for prostate cancer. Eur. Urol. 2011, 59, 823–831. [Google Scholar] [CrossRef] [PubMed]
  66. Valadi, H.; Ekström, K.; Bossios, A.; Sjöstrand, M.; Lee, J.J.; Lötvall, J.O. Exosome-mediated transfer of mRNAs and microRNAs is a novel mechanism of genetic exchange between cells. Nat. Cell Biol. 2007, 9, 654. [Google Scholar] [CrossRef]
  67. Hosseini-Beheshti, E.; Pham, S.; Adomat, H.; Li, N.; Tomlinson Guns, E.S. Exosomes as biomarker enriched microvesicles: Characterization of exosomal proteins derived from a panel of prostate cell lines with distinct AR phenotypes. Mol. Cell Proteom. 2012, 11, 863–885. [Google Scholar] [CrossRef] [PubMed]
  68. Li, Y.; Bahassi, E.M. Biofluid-based circulating tumor molecules as diagnostic tools for use in personalized medicine. J. Mol. Biomark. Diagn. 2013, 5, 157–163. [Google Scholar]
  69. Perkins, G.L.; Slater, E.D.; Sanders, G.K.; Prichard, J.G. Serum tumor markers. Am. Fam. Physician 2003, 68, 1075–1088. [Google Scholar]
  70. Kharmate, G.; Hosseini-Beheshti, E.; Caradec, J.; Chin, M.Y.; Guns, E.S.T. Epidermal growth factor receptor in prostate cancer derived exosomes. PLoS ONE 2016, 11. [Google Scholar]
  71. Khan, S.; Jutzy, J.M.; Valenzuela, M.M.A.; Turay, D.; Aspe, J.R.; Ashok, A.; Mirshahidi, S.; Mercola, D.; Lilly, M.B.; Wall, N.R. Plasma-derived exosomal survivin, a plausible biomarker for early detection of prostate cancer. PLoS ONE 2012, 7, e46737. [Google Scholar] [CrossRef]
  72. Panigrahi, G.K.; Praharaj, P.P.; Kittaka, H.; Mridha, A.R.; Black, O.M.; Singh, R.; Mercer, R.; van Bokhoven, A.; Torkko, K.C.; Agarwal, C. Exosome proteomic analyses identify inflammatory phenotype and novel biomarkers in African American prostate cancer patients. Cancer Med. 2019, 8, 1110–1123. [Google Scholar] [CrossRef] [PubMed]
  73. McKiernan, J.; Donovan, M.J.; O’Neill, V.; Bentink, S.; Noerholm, M.; Belzer, S.; Skog, J.; Kattan, M.W.; Partin, A.; Andriole, G. A novel urine exosome gene expression assay to predict high-grade prostate cancer at initial biopsy. Jama Oncol. 2016, 2, 882–889. [Google Scholar] [CrossRef] [PubMed]
  74. Huang, X.; Yuan, T.; Liang, M.; Du, M.; Xia, S.; Dittmar, R.; Wang, D.; See, W.; Costello, B.A.; Quevedo, F. Exosomal miR-1290 and miR-375 as prognostic markers in castration-resistant prostate cancer. Eur. Urol. 2015, 67, 33–41. [Google Scholar] [CrossRef] [PubMed]
  75. Corcoran, C.; Rani, S.; O’driscoll, L. miR-34a is an intracellular and exosomal predictive biomarker for response to docetaxel with clinical relevance to prostate cancer progression. Prostate 2014, 74, 1320–1334. [Google Scholar] [CrossRef] [PubMed]
  76. Shedden, K.; Xie, X.T.; Chandaroy, P.; Chang, Y.T.; Rosania, G.R. Expulsion of small molecules in vesicles shed by cancer cells: Association with gene expression and chemosensitivity profiles. Cancer Res. 2003, 63, 4331–4337. [Google Scholar] [PubMed]
  77. Safaei, R.; Larson, B.J.; Cheng, T.C.; Gibson, M.A.; Otani, S.; Naerdemann, W.; Howell, S.B. Abnormal lysosomal trafficking and enhanced exosomal export of cisplatin in drug-resistant human ovarian carcinoma cells. Mol. Cancer Ther. 2005, 4, 1595–1604. [Google Scholar] [CrossRef] [PubMed]
  78. Khan, R.; Zahid, S.; Wan, Y.-J.Y.; Forster, J.; Karim, A.-B.A.; Nawabi, A.M.; Azhar, A.; Rahman, M.A.; Ahmed, N. Protein expression profiling of nuclear membrane protein reveals potential biomarker of human hepatocellular carcinoma. Clin. Proteom. 2013, 10, 6. [Google Scholar] [CrossRef]
  79. Aung, T.; Chapuy, B.; Vogel, D.; Wenzel, D.; Oppermann, M.; Lahmann, M.; Weinhage, T.; Menck, K.; Hupfeld, T.; Koch, R. Exosomal evasion of humoral immunotherapy in aggressive B-cell lymphoma modulated by ATP-binding cassette transporter A3. Proc. Natl. Acad. Sci. USA 2011, 108, 15336–15341. [Google Scholar] [CrossRef]
  80. Ciravolo, V.; Huber, V.; Ghedini, G.C.; Venturelli, E.; Bianchi, F.; Campiglio, M.; Morelli, D.; Villa, A.; Mina, P.D.; Menard, S. Potential role of HER2-overexpressing exosomes in countering trastuzumab-based therapy. J. Cell. Physiol. 2012, 227, 658–667. [Google Scholar] [CrossRef]
  81. Zhang, H.-G.; Grizzle, W.E. Exosomes and cancer: A newly described pathway of immune suppression. Clin. Cancer Res. 2011, 17, 959–964. [Google Scholar] [CrossRef]
  82. Alvarez-Erviti, L.; Seow, Y.; Yin, H.; Betts, C.; Lakhal, S.; Wood, M.J. Delivery of siRNA to the mouse brain by systemic injection of targeted exosomes. Nat. Biotechnol. 2011, 29, 341. [Google Scholar] [CrossRef] [PubMed]
  83. Ohno, S.-i.; Ishikawa, A.; Kuroda, M. Roles of exosomes and microvesicles in disease pathogenesis. Adv. Drug Deliv. Rev. 2013, 65, 398–401. [Google Scholar] [CrossRef] [PubMed]
  84. Camussi, G.; Deregibus, M.C.; Bruno, S.; Cantaluppi, V.; Biancone, L. Exosomes/microvesicles as a mechanism of cell-to-cell communication. Kidney Int. 2010, 78, 838–848. [Google Scholar] [CrossRef] [PubMed]
  85. Ramachandran, S.; Palanisamy, V. Horizontal transfer of RNAs: Exosomes as mediators of intercellular communication. Wiley Interdiscip. Rev. 2012, 3, 286–293. [Google Scholar] [CrossRef]
  86. Han, L.; Xu, J.; Xu, Q.; Zhang, B.; Lam, E.F.; Sun, Y. Extracellular vesicles in the tumor microenvironment: Therapeutic resistance, clinical biomarkers, and targeting strategies. Med. Res. Rev. 2017, 37, 1318–1349. [Google Scholar] [CrossRef]
  87. Soekmadji, C.; Riches, J.D.; Russell, P.J.; Ruelcke, J.E.; McPherson, S.; Wang, C.; Hovens, C.M.; Corcoran, N.M.; BioResource, T.A.P.C.C.; Hill, M.M. Modulation of paracrine signaling by CD9 positive small extracellular vesicles mediates cellular growth of androgen deprived prostate cancer. Oncotarget 2017, 8, 52237. [Google Scholar] [CrossRef]
  88. Souza, A.G.; B Silva, I.B.; Campos-Fernández, E.; Marangoni, K.; F Bastos, V.A.; Alves, P.T.; Goulart, L.R.; Alonso-Goulart, V. Extracellular vesicles as drivers of epithelial-mesenchymal transition and carcinogenic characteristics in normal prostate cells. Mol. Carcinog. 2018, 57, 503–511. [Google Scholar] [CrossRef]
  89. Hosseini-Beheshti, E.; Choi, W.; Weiswald, L.-B.; Kharmate, G.; Ghaffari, M.; Roshan-Moniri, M.; Hassona, M.D.; Chan, L.; Chin, M.Y.; Tai, I.T. Exosomes confer pro-survival signals to alter the phenotype of prostate cells in their surrounding environment. Oncotarget 2016, 7, 14639. [Google Scholar] [CrossRef]
  90. Waugh, D.J.; Wilson, C. The interleukin-8 pathway in cancer. Clin. Cancer Res. 2008, 14, 6735–6741. [Google Scholar] [CrossRef]
  91. Ramteke, A.; Ting, H.; Agarwal, C.; Mateen, S.; Somasagara, R.; Hussain, A.; Graner, M.; Frederick, B.; Agarwal, R.; Deep, G. Exosomes secreted under hypoxia enhance invasiveness and stemness of prostate cancer cells by targeting adherens junction molecules. Mol. Carcinog. 2015, 54, 554–565. [Google Scholar] [CrossRef]
  92. Fedele, C.; Singh, A.; Zerlanko, B.J.; Iozzo, R.V.; Languino, L.R. The αvβ6 integrin is transferred intercellularly via exosomes. J. Biol. Chem. 2015, 290, 4545–4551. [Google Scholar] [CrossRef] [PubMed]
  93. Singh, A.; Fedele, C.; Lu, H.; Nevalainen, M.T.; Keen, J.H.; Languino, L.R. Exosome-mediated transfer of αvβ3 integrin from tumorigenic to nontumorigenic cells promotes a migratory phenotype. Mol. Cancer Res. 2016, 14, 1136–1146. [Google Scholar] [CrossRef] [PubMed]
  94. Corcoran, C.; Rani, S.; O’Brien, K.; O’Neill, A.; Prencipe, M.; Sheikh, R.; Webb, G.; McDermott, R.; Watson, W.; Crown, J. Docetaxel-resistance in prostate cancer: Evaluating associated phenotypic changes and potential for resistance transfer via exosomes. PLoS ONE 2012, 7, e50999. [Google Scholar] [CrossRef] [PubMed]
  95. Panagopoulos, K.; Cross-Knorr, S.; Dillard, C.; Pantazatos, D.; Del Tatto, M.; Mills, D.; Goldstein, L.; Renzulli, J.; Quesenberry, P.; Chatterjee, D. Reversal of chemosensitivity and induction of cell malignancy of a non-malignant prostate cancer cell line upon extracellular vesicle exposure. Mol. Cancer 2013, 12, 118. [Google Scholar] [CrossRef] [PubMed]
  96. Wang, J.Q.; DeChalus, A.; Chatterjee, D.N.; Keller, E.T.; Mizokami, A.; Camussi, G.; Mendelsohn, A.R.; Renzulli, J.F. Extracellular vesicle-mediated reversal of paclitaxel resistance in prostate cancer. Crit. Rev. Oncog. 2015, 20, 407. [Google Scholar] [CrossRef]
  97. Tomasetti, M.; Nocchi, L.; Staffolani, S.; Manzella, N.; Amati, M.; Goodwin, J.; Kluckova, K.; Nguyen, M.; Strafella, E.; Bajzikova, M. MicroRNA-126 suppresses mesothelioma malignancy by targeting IRS1 and interfering with the mitochondrial function. Antioxid. Redox Signal. 2014, 21, 2109–2125. [Google Scholar] [CrossRef]
  98. Tomasetti, M.; Monaco, F.; Manzella, N.; Rohlena, J.; Rohlenova, K.; Staffolani, S.; Gaetani, S.; Ciarapica, V.; Amati, M.; Bracci, M. MicroRNA-126 induces autophagy by altering cell metabolism in malignant mesothelioma. Oncotarget 2016, 7, 36338. [Google Scholar] [CrossRef]
  99. Chang, Q.; Li, Y.; White, M.F.; Fletcher, J.A.; Xiao, S. Constitutive activation of insulin receptor substrate 1 is a frequent event in human tumors: Therapeutic implications. Cancer Res. 2002, 62, 6035–6038. [Google Scholar]
  100. Huang, T.; Chu, T. Repression of miR-126 and upregulation of adrenomedullin in the stromal endothelium by cancer-stromal cross talks confers angiogenesis of cervical cancer. Oncogene 2014, 33, 3636. [Google Scholar] [CrossRef]
  101. Sun, X.; Wang, Z.-M.; Song, Y.; Tai, X.-H.; Ji, W.-Y.; Gu, H. MicroRNA-126 modulates the tumor microenvironment by targeting calmodulin-regulated spectrin-associated protein 1 (Camsap1). Int. J. Oncol. 2014, 44, 1678–1684. [Google Scholar] [CrossRef]
  102. Garcia, N.A.; Ontoria-Oviedo, I.; González-King, H.; Diez-Juan, A.; Sepúlveda, P. Glucose starvation in cardiomyocytes enhances exosome secretion and promotes angiogenesis in endothelial cells. PLoS ONE 2015, 10, e0138849. [Google Scholar] [CrossRef] [PubMed]
  103. Garcia, N.A.; Moncayo-Arlandi, J.; Sepulveda, P.; Diez-Juan, A. Cardiomyocyte exosomes regulate glycolytic flux in endothelium by direct transfer of GLUT transporters and glycolytic enzymes. Cardiovasc. Res. 2015, 109, 397–408. [Google Scholar] [CrossRef] [PubMed]
  104. Valis, K.; Prochazka, L.; Boura, E.; Chladova, J.; Obsil, T.; Rohlena, J.; Truksa, J.; Dong, L.-F.; Ralph, S.J.; Neuzil, J. Hippo/Mst1 stimulates transcription of the proapoptotic mediator NOXA in a FoxO1-dependent manner. Cancer Res. 2011, 71, 946–954. [Google Scholar] [CrossRef] [PubMed]
  105. Fong, M.Y.; Zhou, W.; Liu, L.; Alontaga, A.Y.; Chandra, M.; Ashby, J.; Chow, A.; O’Connor, S.T.F.; Li, S.; Chin, A.R. Breast-cancer-secreted miR-122 reprograms glucose metabolism in premetastatic niche to promote metastasis. Nat. Cell Biol. 2015, 17, 183. [Google Scholar] [CrossRef]
  106. Warburg, O. On the origin of cancer cells. Science 1956, 123, 309–314. [Google Scholar] [CrossRef]
  107. Tomasetti, M.; Lee, W.; Santarelli, L.; Neuzil, J. Exosome-derived microRNAs in cancer metabolism: Possible implications in cancer diagnostics and therapy. Exp. Mol. Med. 2017, 49, e285. [Google Scholar] [CrossRef]
  108. Castellana, D.; Zobairi, F.; Martinez, M.C.; Panaro, M.A.; Mitolo, V.; Freyssinet, J.-M.; Kunzelmann, C. Membrane microvesicles as actors in the establishment of a favorable prostatic tumoral niche: A role for activated fibroblasts and CX3CL1-CX3CR1 axis. Cancer Res. 2009, 69, 785–793. [Google Scholar] [CrossRef]
  109. Martinez-Outschoorn, U.E.; Lisanti, M.P.; Sotgia, F. Catabolic cancer-associated fibroblasts transfer energy and biomass to anabolic cancer cells, fueling tumor growth. Semin. Cancer Biol. 2014, 25, 47–60. [Google Scholar] [CrossRef]
  110. Shiga, K.; Hara, M.; Nagasaki, T.; Sato, T.; Takahashi, H.; Takeyama, H. Cancer-associated fibroblasts: Their characteristics and their roles in tumor growth. Cancers 2015, 7, 2443–2458. [Google Scholar] [CrossRef]
  111. Cirri, P.; Chiarugi, P. Cancer-associated-fibroblasts and tumour cells: A diabolic liaison driving cancer progression. Cancer Metastasis Rev. 2012, 31, 195–208. [Google Scholar] [CrossRef]
  112. Flaberg, E.; Markasz, L.; Petranyi, G.; Stuber, G.; Dicső, F.; Alchihabi, N.; Oláh, È.; Csízy, I.; Józsa, T.; Andrén, O. High-throughput live-cell imaging reveals differential inhibition of tumor cell proliferation by human fibroblasts. Int. J. Cancer 2011, 128, 2793–2802. [Google Scholar] [CrossRef] [PubMed]
  113. Öhlund, D.; Elyada, E.; Tuveson, D. Fibroblast heterogeneity in the cancer wound. J. Exp. Med. 2014, 211, 1503–1523. [Google Scholar] [CrossRef] [PubMed]
  114. Fiaschi, T.; Marini, A.; Giannoni, E.; Taddei, M.L.; Gandellini, P.; De Donatis, A.; Lanciotti, M.; Serni, S.; Cirri, P.; Chiarugi, P. Reciprocal metabolic reprogramming through lactate shuttle coordinately influences tumor-stroma interplay. Cancer Res. 2012, 72, 5130–5140. [Google Scholar] [CrossRef] [PubMed]
  115. Vander Heiden, M.G.; Cantley, L.C.; Thompson, C.B. Understanding the Warburg effect: The metabolic requirements of cell proliferation. Science 2009, 324, 1029–1033. [Google Scholar] [CrossRef]
  116. Zhao, H.; Yang, L.; Baddour, J.; Achreja, A.; Bernard, V.; Moss, T.; Marini, J.C.; Tudawe, T.; Seviour, E.G.; San Lucas, F.A. Tumor microenvironment derived exosomes pleiotropically modulate cancer cell metabolism. Elife 2016, 5, e10250. [Google Scholar] [CrossRef]
  117. Paggetti, J.; Haderk, F.; Seiffert, M.; Janji, B.; Distler, U.; Ammerlaan, W.; Kim, Y.J.; Adam, J.; Lichter, P.; Solary, E. Exosomes released by chronic lymphocytic leukemia cells induce the transition of stromal cells into cancer-associated fibroblasts. Blood 2015, 126, 1106–1117. [Google Scholar] [CrossRef]
  118. Quail, D.F.; Joyce, J.A. Microenvironmental regulation of tumor progression and metastasis. Nat. Med. 2013, 19, 1423. [Google Scholar] [CrossRef]
  119. Duijvesz, D.; Burnum-Johnson, K.E.; Gritsenko, M.A.; Hoogland, A.M.; Vredenbregt-van den Berg, M.S.; Willemsen, R.; Luider, T.; Paša-Tolić, L.; Jenster, G. Proteomic profiling of exosomes leads to the identification of novel biomarkers for prostate cancer. PLoS ONE 2013, 8, e82589. [Google Scholar] [CrossRef]
  120. Ronquist, K.G.; Ronquist, G.; Larsson, A.; Carlsson, L. Proteomic analysis of prostate cancer metastasis-derived prostasomes. Anticancer Res. 2010, 30, 285–290. [Google Scholar]
  121. Bang, C.; Thum, T. Exosomes: New players in cell–cell communication. Int. J. Biochem. Cell Biol. 2012, 44, 2060–2064. [Google Scholar] [CrossRef]
  122. Mathivanan, S.; Ji, H.; Simpson, R.J. Exosomes: Extracellular organelles important in intercellular communication. J. Proteom. 2010, 73, 1907–1920. [Google Scholar] [CrossRef] [PubMed]
  123. Raposo, G.; Stoorvogel, W. Extracellular vesicles: Exosomes, microvesicles, and friends. J. Cell Biol. 2013, 200, 373–383. [Google Scholar] [CrossRef] [PubMed]
  124. Beach, A.; Zhang, H.-G.; Ratajczak, M.Z.; Kakar, S.S. Exosomes: An overview of biogenesis, composition and role in ovarian cancer. J. Ovarian Res. 2014, 7, 14. [Google Scholar] [CrossRef] [PubMed]
  125. Hannafon, B.; Ding, W.-Q. Intercellular communication by exosome-derived microRNAs in cancer. Int. J. Mol. Sci. 2013, 14, 14240–14269. [Google Scholar] [CrossRef] [PubMed]
  126. Vlassov, A.V.; Magdaleno, S.; Setterquist, R.; Conrad, R. Exosomes: Current knowledge of their composition, biological functions, and diagnostic and therapeutic potentials. Biochim. Biophys. Acta 2012, 1820, 940–948. [Google Scholar] [CrossRef] [PubMed]
  127. Harshman, S.W.; Canella, A.; Ciarlariello, P.D.; Rocci, A.; Agarwal, K.; Smith, E.M.; Talabere, T.; Efebera, Y.A.; Hofmeister, C.C.; Benson, D.M., Jr. Characterization of multiple myeloma vesicles by label-free relative quantitation. Proteomics 2013, 13, 3013–3029. [Google Scholar] [CrossRef]
  128. Gonzalez-Begne, M.; Lu, B.; Liao, L.; Xu, T.; Bedi, G.; Melvin, J.E.; Yates, J.R., III. Characterization of the human submandibular/sublingual saliva glycoproteome using lectin affinity chromatography coupled to multidimensional protein identification technology. J. Proteome Res. 2011, 10, 5031–5046. [Google Scholar] [CrossRef]
  129. Gonzalez-Begne, M.; Lu, B.; Han, X.; Hagen, F.K.; Hand, A.R.; Melvin, J.E.; Yates, J.R., III. Proteomic analysis of human parotid gland exosomes by multidimensional protein identification technology (MudPIT). J. Proteome Res. 2009, 8, 1304–1314. [Google Scholar] [CrossRef]
  130. Abd Elmageed, Z.Y.; Yang, Y.; Thomas, R.; Ranjan, M.; Mondal, D.; Moroz, K.; Fang, Z.; Rezk, B.M.; Moparty, K.; Sikka, S.C. Neoplastic reprogramming of patient-derived adipose stem cells by prostate cancer cell-associated exosomes. Stem Cells 2014, 32, 983–997. [Google Scholar] [CrossRef]
  131. Webber, J.; Steadman, R.; Mason, M.D.; Tabi, Z.; Clayton, A. Cancer exosomes trigger fibroblast to myofibroblast differentiation. Cancer Res. 2010, 70, 9621–9630. [Google Scholar] [CrossRef]
  132. Webber, J.P.; Spary, L.K.; Sanders, A.J.; Chowdhury, R.; Jiang, W.G.; Steadman, R.; Wymant, J.; Jones, A.T.; Kynaston, H.; Mason, M.D. Differentiation of tumour-promoting stromal myofibroblasts by cancer exosomes. Oncogene 2015, 34, 290. [Google Scholar] [CrossRef] [PubMed]
  133. Tomasek, J.J.; Gabbiani, G.; Hinz, B.; Chaponnier, C.; Brown, R.A. Myofibroblasts and mechano-regulation of connective tissue remodelling. Nat. Rev. Mol. Cell Biol. 2002, 3, 349. [Google Scholar] [CrossRef] [PubMed]
  134. Calvo, F.; Ege, N.; Grande-Garcia, A.; Hooper, S.; Jenkins, R.P.; Chaudhry, S.I.; Harrington, K.; Williamson, P.; Moeendarbary, E.; Charras, G. Mechanotransduction and YAP-dependent matrix remodelling is required for the generation and maintenance of cancer-associated fibroblasts. Nat. Cell Biol. 2013, 15, 637. [Google Scholar] [CrossRef] [PubMed]
  135. Shimoda, M.; Principe, S.; Jackson, H.W.; Luga, V.; Fang, H.; Molyneux, S.D.; Shao, Y.W.; Aiken, A.; Waterhouse, P.D.; Karamboulas, C. Loss of the Timp gene family is sufficient for the acquisition of the CAF-like cell state. Nat. Cell Biol. 2014, 16, 889. [Google Scholar] [CrossRef]
  136. Chowdhury, R.; Webber, J.P.; Gurney, M.; Mason, M.D.; Tabi, Z.; Clayton, A. Cancer exosomes trigger mesenchymal stem cell differentiation into pro-angiogenic and pro-invasive myofibroblasts. Oncotarget 2015, 6, 715. [Google Scholar] [CrossRef]
  137. Hoshino, A.; Costa-Silva, B.; Shen, T.-L.; Rodrigues, G.; Hashimoto, A.; Mark, M.T.; Molina, H.; Kohsaka, S.; Di Giannatale, A.; Ceder, S. Tumour exosome integrins determine organotropic metastasis. Nature 2015, 527, 329. [Google Scholar] [CrossRef]
  138. Tatarov, O.; Mitchell, T.J.; Seywright, M.; Leung, H.Y.; Brunton, V.G.; Edwards, J. SRC family kinase activity is up-regulated in hormone-refractory prostate cancer. Clin. Cancer Res. 2009, 15, 3540–3549. [Google Scholar] [CrossRef]
  139. Chang, C.Y.-M.; Kung, H.-J.; Evans, C.P. Nonreceptor Tyrosine Kinases in Prostate. Neoplasia 2007, 9, 90–100. [Google Scholar] [CrossRef]
  140. Marx, M.; Warren, S.L.; Madri, J.A. pp60c-src modulates microvascular endothelial phenotype and in vitro angiogenesis. Exp. Mol. Pathol. 2001, 70, 201–213. [Google Scholar] [CrossRef]
  141. DeRita, R.M.; Zerlanko, B.; Singh, A.; Lu, H.; Iozzo, R.V.; Benovic, J.L.; Languino, L.R. c-Src, insulin-like growth factor I receptor, G-protein-coupled receptor kinases and focal adhesion kinase are enriched into prostate cancer cell exosomes. J. Cell. Biochem. 2017, 118, 66–73. [Google Scholar] [CrossRef]
  142. Angelucci, A.; D’ascenzo, S.; Festuccia, C.; Gravina, G.L.; Bologna, M.; Dolo, V.; Pavan, A. Vesicle-associated urokinase plasminogen activator promotes invasion in prostate cancer cell lines. Clin. Exp. Metastasis 2000, 18, 163. [Google Scholar] [CrossRef] [PubMed]
  143. Drew, A.F.; Tucker, H.L.; Kombrinck, K.W.; Simon, D.I.; Bugge, T.H.; Degen, J.L. Plasminogen is a critical determinant of vascular remodeling in mice. Circ. Res. 2000, 87, 133–139. [Google Scholar] [CrossRef] [PubMed]
  144. Lundholm, M.; Schröder, M.; Nagaeva, O.; Baranov, V.; Widmark, A.; Mincheva-Nilsson, L.; Wikström, P. Prostate tumor-derived exosomes down-regulate NKG2D expression on natural killer cells and CD8+ T cells: Mechanism of immune evasion. PLoS ONE 2014, 9, e108925. [Google Scholar] [CrossRef] [PubMed]
  145. Naito, Y.; Yoshioka, Y.; Yamamoto, Y.; Ochiya, T. How cancer cells dictate their microenvironment: Present roles of extracellular vesicles. Cell. Mol. Life Sci. 2017, 74, 697–713. [Google Scholar] [CrossRef]
  146. Steinbichler, T.B.; Dudas, J.; Riechelmann, H.; Skvortsova, I.-I. The role of exosomes in cancer metastasis. Semin. Cancer Biol. 2017, 44, 170–181. [Google Scholar] [CrossRef]
  147. Meehan, B.; Rak, J.; Di Vizio, D. Oncosomes–large and small: What are they, where they came from? J. Extracell. Vesicles 2016, 5. [Google Scholar] [CrossRef]
  148. Minciacchi, V.R.; Spinelli, C.; Reis-Sobreiro, M.; Cavallini, L.; You, S.; Zandian, M.; Li, X.; Mishra, R.; Chiarugi, P.; Adam, R.M. MYC mediates large oncosome-induced fibroblast reprogramming in prostate cancer. Cancer Res. 2017, 77, 2306–2317. [Google Scholar] [CrossRef]
  149. Silva, T.A.; Smuczek, B.; Valadao, I.C.; Dzik, L.M.; Iglesia, R.P.; Cruz, M.C.; Zelanis, A.; de Siqueira, A.S.; Serrano, S.M.; Goldberg, G.S.; et al. AHNAK enables mammary carcinoma cells to produce extracellular vesicles that increase neighboring fibroblast cell motility. Oncotarget 2016, 7, 49998–50016. [Google Scholar] [CrossRef]
  150. Mizutani, K.; Terazawa, R.; Kameyama, K.; Kato, T.; Horie, K.; Tsuchiya, T.; Seike, K.; Ehara, H.; Fujita, Y.; Kawakami, K. Isolation of prostate cancer-related exosomes. Anticancer Res. 2014, 34, 3419–3423. [Google Scholar]
  151. Lehmann, B.D.; Paine, M.S.; Brooks, A.M.; McCubrey, J.A.; Renegar, R.H.; Wang, R.; Terrian, D.M. Senescence-associated exosome release from human prostate cancer cells. Cancer Res. 2008, 68, 7864–7871. [Google Scholar] [CrossRef]
  152. Itoh, T.; Ito, Y.; Ohtsuki, Y.; Tsukamasa, Y.; Yamada, N.; Naoe, T.; Akao, Y. Microvesicles released from hormone-refractory prostate cancer cells facilitate mouse pre-osteoblast differentiation. J. Mol. Histol. 2012, 43, 509–515. [Google Scholar] [CrossRef] [PubMed]
  153. Laderach, D.J.; Gentilini, L.D.; Giribaldi, L.; Delgado, V.C.; Nugnes, L.; Croci, D.O.; Al Nakouzi, N.; Sacca, P.; Casas, G.; Mazza, O. A unique galectin signature in human prostate cancer progression suggests galectin-1 as a key target for treatment of advanced disease. Cancer Res. 2013, 73, 86–96. [Google Scholar] [CrossRef] [PubMed]
  154. Kawakami, K.; Fuhita, Y.; Kato, T.; Mizutani, K.; Kameyama, K.; Tsumoto, H.; Miura, Y.; Deguchi, T.; Ito, M. Integrin beta4 and vinculin contained in exosomes are potential markers for progression of prostate cancer associated with taxane-resistance. Int. J. Oncol 2015, 47, 384–390. [Google Scholar] [CrossRef] [PubMed]
  155. Kato, T.; Mizutani, K.; Kameyama, K.; Kawakami, K.; Fujita, Y.; Nakane, K.; Kanimoto, Y.; Ehara, H.; Ito, H.; Seishima, M. Serum exosomal P-glycoprotein is a potential marker to diagnose docetaxel resistance and select a taxoid for patients with prostate cancer. Urol. Oncol. 2015, 33, 385. [Google Scholar] [CrossRef] [PubMed]
  156. Wang, X.; Wilson, M.J.; Slaton, J.W.; Sinha, A.A.; Ewing, S.L.; Pei, D. Increased Aggressiveness of Human Prostate PC-3 Tumor Cells Expressing Cell Surface Localized Membrane Type-1 Matrix Metalloproteinase (MT1-MMP). J. Androl. 2009, 30, 259–274. [Google Scholar] [CrossRef] [PubMed]
  157. Chen, G.; Huang, A.C.; Zhang, W.; Zhang, G.; Wu, M.; Xu, W.; Yu, Z.; Yang, J.; Wang, B.; Sun, H.; et al. Exosomal PD-L1 contributes to immunosuppression and is associated with anti-PD-1 response. Nature 2018, 560, 382–386. [Google Scholar] [CrossRef] [PubMed]
  158. Oh, W.K.; Vargas, R.; Jacobus, S.; Leitzel, K.; Regan, M.M.; Hamer, P.; Pierce, K.; Brown-Shimer, S.; Carney, W.; Ali, S.M. Elevated plasma tissue inhibitor of metalloproteinase-1 levels predict decreased survival in castration-resistant prostate cancer patients. Cancer 2011, 117, 517–525. [Google Scholar] [CrossRef]
  159. Trerotola, M.; Ganguly, K.K.; Fazli, L.; Fedele, C.; Lu, H.; Dutta, A.; Liu, Q.; De Angelis, T.; Riddell, L.W.; Riobo, N.A. Trop-2 is up-regulated in invasive prostate cancer and displaces FAK from focal contacts. Oncotarget 2015, 6, 14318. [Google Scholar] [CrossRef]
  160. El-Sayed, I.Y.; Daher, A.; Destouches, D.; Firlej, V.; Kostallari, E.; Maillé, P.; Huet, E.; Haidar-Ahmad, N.; Jenster, G.; de la Taille, A. Extracellular vesicles released by mesenchymal-like prostate carcinoma cells modulate EMT state of recipient epithelial-like carcinoma cells through regulation of AR signaling. Cancer Lett. 2017, 410, 100–111. [Google Scholar] [CrossRef]
  161. Charrin, S.; Manié, S.; Thiele, C.; Billard, M.; Gerlier, D.; Boucheix, C.; Rubinstein, E. A physical and functional link between cholesterol and tetraspanins. Eur. J. Immunol. 2003, 33, 2479–2489. [Google Scholar] [CrossRef]
  162. Record, M.; Carayon, K.; Poirot, M.; Silvente-Poirot, S. Exosomes as new vesicular lipid transporters involved in cell–cell communication and various pathophysiologies. Biochim. Biophys. Acta 2014, 1841, 108–120. [Google Scholar] [CrossRef] [PubMed]
  163. Zakharova, L.; Svetlova, M.; Fomina, A.F. T cell exosomes induce cholesterol accumulation in human monocytes via phosphatidylserine receptor. J. Cell. Physiol. 2007, 212, 174–181. [Google Scholar] [CrossRef]
  164. Wang, D.; DuBois, R.N. Eicosanoids and cancer. Nat. Rev. Cancer 2010, 10, 181. [Google Scholar] [CrossRef] [PubMed]
  165. Beloribi, S.; Ristorcelli, E.; Breuzard, G.; Silvy, F.; Bertrand-Michel, J.; Beraud, E.; Verine, A.; Lombardo, D. Exosomal lipids impact notch signaling and induce death of human pancreatic tumoral SOJ-6 cells. PLoS ONE 2012, 7, e47480. [Google Scholar] [CrossRef]
  166. Beloribi-Djefaflia, S.; Siret, C.; Lombardo, D. Exosomal lipids induce human pancreatic tumoral MiaPaCa-2 cells resistance through the CXCR4-SDF-1α signaling axis. Oncoscience 2015, 2, 15. [Google Scholar] [PubMed]
  167. Llorente, A.; Skotland, T.; Sylvänne, T.; Kauhanen, D.; Róg, T.; Orłowski, A.; Vattulainen, I.; Ekroos, K.; Sandvig, K. Molecular lipidomics of exosomes released by PC-3 prostate cancer cells. Biochim. Biophys. Acta 2013, 1831, 1302–1309. [Google Scholar] [CrossRef]
  168. Prantl, L.; Muehlberg, F.; Navone, N.M.; Song, Y.H.; Vykoukal, J.; Logothetis, C.J.; Alt, E.U. Adipose tissue-derived stem cells promote prostate tumor growth. Prostate 2010, 70, 1709–1715. [Google Scholar] [CrossRef]
  169. Gajos-Michniewicz, A.; Duechler, M.; Czyz, M. MiRNA in melanoma-derived exosomes. Cancer Lett. 2014, 347, 29–37. [Google Scholar] [CrossRef]
  170. Bartel, D.P. MicroRNAs: Target recognition and regulatory functions. Cell 2009, 136, 215–233. [Google Scholar] [CrossRef]
  171. Lim, L.P.; Lau, N.C.; Garrett-Engele, P.; Grimson, A.; Schelter, J.M.; Castle, J.; Bartel, D.P.; Linsley, P.S.; Johnson, J.M. Microarray analysis shows that some microRNAs downregulate large numbers of target mRNAs. Nature 2005, 433, 769–773. [Google Scholar] [CrossRef]
  172. Song, J.J.; Smith, S.K.; Hannon, G.J.; Joshua-Tor, L. Crystal structure of Argonaute and its implications for RISC slicer activity. Science 2004, 305, 1434–1437. [Google Scholar] [CrossRef] [PubMed]
  173. Jiang, X.; Hu, S.; Liu, Q.; Qian, C.; Liu, Z.; Luo, D. Exosomal microRNA remodels the tumor microenvironment. Peer J. 2017, 5, e4196. [Google Scholar] [CrossRef] [PubMed]
  174. Pang, W.; Su, J.; Wang, Y.; Feng, H.; Dai, X.; Yuan, Y.; Chen, X.; Yao, W. Pancreatic cancer-secreted miR-155 implicates in the conversion from normal fibroblasts to cancer-associated fibroblasts. Cancer Sci. 2015, 106, 1362–1369. [Google Scholar] [CrossRef] [PubMed]
  175. Renzulli, J.F., II; Del Tatto, M.; Dooner, G.; Aliotta, J.; Goldstein, L.; Dooner, M.; Colvin, G.; Chatterjee, D.; Quesenberry, P. Microvesicle induction of prostate specific gene expression in normal human bone marrow cells. J. Urol. 2010, 184, 2165–2171. [Google Scholar] [CrossRef]
  176. Sánchez, C.A.; Andahur, E.I.; Valenzuela, R.; Castellon, E.A.; Fulla, J.A.; Ramos, C.G.; Trivino, J.C. Exosomes from bulk and stem cells from human prostate cancer have a differential microRNA content that contributes cooperatively over local and pre-metastatic niche. Oncotarget 2016, 7, 3993. [Google Scholar] [CrossRef] [PubMed]
  177. Shiao, S.L.; Chu, G.C.-Y.; Chung, L.W. Regulation of prostate cancer progression by the tumor microenvironment. Cancer Lett. 2016, 380, 340–348. [Google Scholar] [CrossRef]
  178. Chu, G.C.-Y.; Zhau, H.E.; Wang, R.; Rogatko, A.; Feng, X.; Zayzafoon, M.; Liu, Y.; Farach-Carson, M.C.; You, S.; Kim, J. RANK-and c-Met-mediated signal network promotes prostate cancer metastatic colonization. Endocr. Relat. Cancer 2014, 21, 311. [Google Scholar] [CrossRef]
  179. Li, Q.; Li, Q.; Nuccio, J.; Liu, C.; Duan, P.; Wang, R.; Jones, L.W.; Chung, L.W.; Zhau, H.E. Metastasis initiating cells in primary prostate cancer tissues from transurethral resection of the prostate (TURP) predicts castration-resistant progression and survival of prostate cancer patients. Prostate 2015, 75, 1312–1321. [Google Scholar] [CrossRef]
  180. Sugatani, T.; Vacher, J.; Hruska, K.A. A microRNA expression signature of osteoclastogenesis. Blood 2011, 117, 3648–3657. [Google Scholar] [CrossRef]
  181. Zhang, H.-L.; Qin, X.-J.; Cao, D.-L.; Zhu, Y.; Yao, X.-D.; Zhang, S.-L.; Dai, B.; Ye, D.-W. An elevated serum miR-141 level in patients with bone-metastatic prostate cancer is correlated with more bone lesions. Asian J. Androl. 2013, 15, 231. [Google Scholar] [CrossRef]
  182. Morhayim, J.; van de Peppel, J.; Demmers, J.A.; Kocer, G.; Nigg, A.L.; van Driel, M.; Chiba, H.; van Leeuwen, J.P. Proteomic signatures of extracellular vesicles secreted by nonmineralizing and mineralizing human osteoblasts and stimulation of tumor cell growth. Faseb J. 2014, 29, 274–285. [Google Scholar] [CrossRef] [PubMed]
  183. Dragomir, M.; Chen, B.; Calin, G.A. Exosomal lncRNAs as new players in cell-to-cell communication. Transl. Cancer Res. 2018, 7, S243. [Google Scholar] [CrossRef] [PubMed]
  184. Ahadi, A.; Brennan, S.; Kennedy, P.J.; Hutvagner, G.; Tran, N. Long non-coding RNAs harboring miRNA seed regions are enriched in prostate cancer exosomes. Sci. Rep. 2016, 6, 24922. [Google Scholar] [CrossRef] [PubMed]
  185. Wang, J.; Yang, X.; Li, R.; Wang, L.; Gu, Y.; Zhao, Y.; Huang, K.H.; Cheng, T.; Yuan, Y.; Gao, S. Long non-coding RNA MYU promotes prostate cancer proliferation by mediating the miR-184/c-Myc axis. Oncol. Rep. 2018, 40, 2814–2825. [Google Scholar] [CrossRef] [PubMed]
  186. Işın, M.; Uysaler, E.; Özgür, E.; Köseoğlu, H.; Şanlı, Ö.; Yücel, Ö.B.; Gezer, U.; Dalay, N. Exosomal lncRNA-p21 levels may help to distinguish prostate cancer from benign disease. Front. Genet. 2015, 6, 168. [Google Scholar]
  187. Kim, J.; Morley, S.; Le, M.; Bedoret, D.; Umetsu, D.T.; Di Vizio, D.; Freeman, M.R. Enhanced shedding of extracellular vesicles from amoeboid prostate cancer cells: Potential effects on the tumor microenvironment. Cancer Biol. Ther. 2014, 15, 409–418. [Google Scholar] [CrossRef]
  188. Liu, C.M.; Hsieh, C.L.; Shen, C.N.; Lin, C.C.; Shigemura, K.; Sung, S.Y. Exosomes from the tumor microenvironment as reciprocal regulators that enhance prostate cancer progression. Int. J. Urol. 2016, 23, 734–744. [Google Scholar] [CrossRef]
  189. Li, S.L.; An, N.; Liu, B.; Wang, S.Y.; Wang, J.J.; Ye, Y. Exosomes from LNCaP cells promote osteoblast activity through miR-375 transfer. Oncol. Lett. 2019, 17, 4463–4473. [Google Scholar] [CrossRef]
  190. Lucotti, S.; Rainaldi, G.; Evangelista, M.; Rizzo, M. Fludarabine treatment favors the retention of miR-485-3p by prostate cancer cells: Implications for survival. Mol. Cancer 2013, 12, 52. [Google Scholar] [CrossRef]
  191. Olumi, A.F.; Grossfeld, G.D.; Hayward, S.W.; Carroll, P.R.; Tlsty, T.D.; Cunha, G.R. Carcinoma-associated fibroblasts direct tumor progression of initiated human prostatic epithelium. Cancer Res. 1999, 59, 5002–5011. [Google Scholar]
  192. Harper, J.; Sainson, R.C. Regulation of the anti-tumour immune response by cancer-associated fibroblasts. Semin. Cancer Biol. 2014, 25, 69–77. [Google Scholar] [CrossRef]
  193. Bhowmick, N.A.; Chytil, A.; Plieth, D.; Gorska, A.E.; Dumont, N.; Shappell, S.; Washington, M.K.; Neilson, E.G.; Moses, H.L. TGF-beta signaling in fibroblasts modulates the oncogenic potential of adjacent epithelia. Science 2004, 303, 848–851. [Google Scholar] [CrossRef]
  194. Bruzzese, F.; Hägglöf, C.; Leone, A.; Sjöberg, E.; Roca, M.S.; Kiflemariam, S.; Sjöblom, T.; Hammarsten, P.; Egevad, L.; Bergh, A.; et al. Local and systemic protumorigenic effects of cancer-associated fibroblast-derived GDF15. Cancer Res. 2014, 74, 3408–3417. [Google Scholar] [CrossRef] [PubMed]
  195. Hayward, S.W.; Wang, Y.; Cao, M.; Hom, Y.K.; Zhang, B.; Grossfeld, G.D.; Sudilovsky, D.; Cunha, G.R. Malignant transformation in a nontumorigenic human prostatic epithelial cell line. Cancer Res. 2001, 61, 8135–8142. [Google Scholar] [PubMed]
  196. Josson, S.; Gururajan, M.; Sung, S.; Hu, P.; Shao, C.; Zhau, H.; Liu, C.; Lichterman, J.; Duan, P.; Li, Q. Stromal fibroblast-derived miR-409 promotes epithelial-to-mesenchymal transition and prostate tumorigenesis. Oncogene 2015, 34, 2690. [Google Scholar] [CrossRef]
  197. Josson, S.; Gururajan, M.; Hu, P.; Shao, C.; Chu, G.C.-Y.; Zhau, H.E.; Liu, C.; Lao, K.; Lu, C.-L.; Lu, Y.-T. miR-409-3p/-5p promotes tumorigenesis, epithelial-to-mesenchymal transition, and bone metastasis of human prostate cancer. Clin. Cancer Res. 2014, 20, 4636–4646. [Google Scholar] [CrossRef]
  198. Yeung, C.L.A.; Co, N.-N.; Tsuruga, T.; Yeung, T.-L.; Kwan, S.-Y.; Leung, C.S.; Li, Y.; Lu, E.S.; Kwan, K.; Wong, K.-K. Exosomal transfer of stroma-derived miR21 confers paclitaxel resistance in ovarian cancer cells through targeting APAF1. Nat. Commun. 2016, 7, 11150. [Google Scholar] [CrossRef]
  199. Alcayaga-Miranda, F.; González, P.L.; Lopez-Verrilli, A.; Varas-Godoy, M.; Aguila-Díaz, C.; Contreras, L.; Khoury, M. Prostate tumor-induced angiogenesis is blocked by exosomes derived from menstrual stem cells through the inhibition of reactive oxygen species. Oncotarget 2016, 7, 44462. [Google Scholar] [CrossRef]
  200. De Visser, K.E.; Eichten, A.; Coussens, L.M. Paradoxical roles of the immune system during cancer development. Nat. Rev. Cancer 2006, 6, 24–37. [Google Scholar] [CrossRef]
  201. Eichmüller, S.B.; Osen, W.; Mandelboim, O.; Seliger, B. Immune modulatory microRNAs involved in tumor attack and tumor immune escape. J. Natl. Cancer Inst. 2017, 109. [Google Scholar]
  202. Fridlender, Z.G.; Sun, J.; Kim, S.; Kapoor, V.; Cheng, G.; Ling, L.; Worthen, G.S.; Albelda, S.M. Polarization of tumor-associated neutrophil phenotype by TGF-β: “N1” versus “N2” TAN. Cancer Cell 2009, 16, 183–194. [Google Scholar] [CrossRef] [PubMed]
  203. Grivennikov, S.I.; Greten, F.R.; Karin, M. Immunity, inflammation, and cancer. Cell 2010, 140, 883–899. [Google Scholar] [CrossRef] [PubMed]
  204. Psaila, B.; Lyden, D. The metastatic niche: Adapting the foreign soil. Nat. Rev. Cancer 2009, 9, 285–293. [Google Scholar] [CrossRef] [PubMed]
  205. Gabrilovich, D.I.; Nagaraj, S. Myeloid-derived suppressor cells as regulators of the immune system. Nat. Rev. Immunol. 2009, 9, 162–174. [Google Scholar] [CrossRef]
  206. Alderson, M.R.; Tough, T.W.; Davis-Smith, T.; Braddy, S.; Falk, B.; Schooley, K.A.; Goodwin, R.G.; Smith, C.; Ramsdell, F.; Lynch, D.H. Fas ligand mediates activation-induced cell death in human T lymphocytes. J. Exp. Med. 1995, 181, 71–77. [Google Scholar] [CrossRef]
  207. Van Parijs, L.; Abbas, A.K. Role of Fas-mediated cell death in the regulation of immune responses. Curr. Opin. Immunol. 1996, 8, 355–361. [Google Scholar] [CrossRef]
  208. Boyiadzis, M.; Whiteside, T. The emerging roles of tumor-derived exosomes in hematological malignancies. Leukemia 2017, 31, 1259–1268. [Google Scholar] [CrossRef]
  209. Filipazzi, P.; Bürdek, M.; Villa, A.; Rivoltini, L.; Huber, V. Recent advances on the role of tumor exosomes in immunosuppression and disease progression. Semin. Cancer Biol. 2012, 22, 342–349. [Google Scholar] [CrossRef]
  210. Whiteside, T.L. Exosomes and tumor-mediated immune suppression. J. Clin. Investig. 2016, 126, 1216–1223. [Google Scholar] [CrossRef]
  211. Chow, A.; Zhou, W.; Liu, L.; Fong, M.Y.; Champer, J.; Van Haute, D.; Chin, A.R.; Ren, X.; Gugiu, B.G.; Meng, Z. Macrophage immunomodulation by breast cancer-derived exosomes requires Toll-like receptor 2-mediated activation of NF-κB. Sci. Rep. 2014, 4, 1–11. [Google Scholar] [CrossRef]
  212. Marton, A.; Vizler, C.; Kusz, E.; Temesfoi, V.; Szathmary, Z.; Nagy, K.; Szegletes, Z.; Varo, G.; Siklos, L.; Katona, R.L. Melanoma cell-derived exosomes alter macrophage and dendritic cell functions in vitro. Immunol. Lett. 2012, 148, 34–38. [Google Scholar] [CrossRef]
  213. Sonda, N.; Simonato, F.; Peranzoni, E.; Calì, B.; Bortoluzzi, S.; Bisognin, A.; Wang, E.; Marincola, F.M.; Naldini, L.; Gentner, B. miR-142-3p prevents macrophage differentiation during cancer-induced myelopoiesis. Immunity 2013, 38, 1236–1249. [Google Scholar] [CrossRef]
  214. Bell, E.; Taylor, M.A. Functional roles for exosomal microRNAs in the tumour microenvironment. Comput. Struct. Biotechnol. J. 2017, 15, 8–13. [Google Scholar] [CrossRef] [PubMed]
  215. Ye, S.B.; Li, Z.L.; Luo, D.H.; Huang, B.J.; Chen, Y.S.; Zhang, X.S.; Cui, J.; Zeng, Y.X.; Li, J. Tumor-derived exosomes promote tumor progression and T-cell dysfunction through the regulation of enriched exosomal microRNAs in human nasopharyngeal carcinoma. Oncotarget 2014, 5, 5439. [Google Scholar] [CrossRef] [PubMed]
  216. Ding, G.; Zhou, L.; Qian, Y.; Fu, M.; Chen, J.; Chen, J.; Xiang, J.; Wu, Z.; Jiang, G.; Cao, L. Pancreatic cancer-derived exosomes transfer miRNAs to dendritic cells and inhibit RFXAP expression via miR-212-3p. Oncotarget 2015, 6, 29877. [Google Scholar] [CrossRef]
  217. Zhou, M.; Chen, J.; Zhou, L.; Chen, W.; Ding, G.; Cao, L. Pancreatic cancer derived exosomes regulate the expression of TLR4 in dendritic cells via miR-203. Cell. Immunol. 2014, 292, 65–69. [Google Scholar] [CrossRef]
  218. Rivoltini, L.; Chiodoni, C.; Squarcina, P.; Tortoreto, M.; Villa, A.; Vergani, B.; Burdek, M.; Botti, L.; Arioli, I.; Cova, A.; et al. TNF-Related Apoptosis-Inducing Ligand (TRAIL)-Armed Exosomes Deliver Proapoptotic Signals to Tumor Site. Clin. Cancer Res. 2016, 22, 3499–3512. [Google Scholar] [CrossRef] [PubMed]
  219. Catalano, V.; Turdo, A.; Di Franco, S.; Dieli, F.; Todaro, M.; Stassi, G. Tumor and its Microenvironment: A Synergistic Interplay. Semin. Cancer Biol. 2013, 23, 522–532. [Google Scholar]
  220. Roma-Rodrigues, C.; Fernandes, A.R.; Baptista, P.V. Exosome in tumour microenvironment: Overview of the crosstalk between normal and cancer cells. Biomed. Res. Int. 2014, 2014. [Google Scholar] [CrossRef]
  221. Ruivo, C.F.; Adem, B.; Silva, M.; Melo, S.A. The biology of cancer exosomes: Insights and new perspectives. Cancer Res. 2017, 77, 6480–6488. [Google Scholar] [CrossRef]
  222. Busby, J.; McMenamin, Ú.; Spence, A.; Johnston, B.; Hughes, C.; Cardwell, C. Angiotensin receptor blocker use and gastro-oesophageal cancer survival: A population-based cohort study. Aliment. Pharmacol. Ther. 2018, 47, 279–288. [Google Scholar] [CrossRef] [PubMed]
  223. Coulson, R.; Liew, S.H.; Connelly, A.A.; Yee, N.S.; Deb, S.; Kumar, B.; Vargas, A.C.; O’Toole, S.A.; Parslow, A.C.; Poh, A. The angiotensin receptor blocker, Losartan, inhibits mammary tumor development and progression to invasive carcinoma. Oncotarget 2017, 8, 18640. [Google Scholar] [CrossRef] [PubMed]
  224. Diop-Frimpong, B.; Chauhan, V.P.; Krane, S.; Boucher, Y.; Jain, R.K. Losartan inhibits collagen I synthesis and improves the distribution and efficacy of nanotherapeutics in tumors. Proc. Natl. Acad. Sci. USA 2011, 108, 2909–2914. [Google Scholar] [CrossRef] [PubMed]
  225. Cassinelli, G.; Lanzi, C.; Tortoreto, M.; Cominetti, D.; Petrangolini, G.; Favini, E.; Zaffaroni, N.; Pisano, C.; Penco, S.; Vlodavsky, I. Antitumor efficacy of the heparanase inhibitor SST0001 alone and in combination with antiangiogenic agents in the treatment of human pediatric sarcoma models. Biochem. Pharmacol. 2013, 85, 1424–1432. [Google Scholar] [CrossRef] [PubMed]
  226. Ritchie, J.P.; Ramani, V.C.; Ren, Y.; Naggi, A.; Torri, G.; Casu, B.; Penco, S.; Pisano, C.; Carminati, P.; Tortoreto, M. SST0001, a chemically modified heparin, inhibits myeloma growth and angiogenesis via disruption of the heparanase/syndecan-1 axis. Clin. Cancer Res. 2011, 17, 1382–1393. [Google Scholar] [CrossRef]
  227. Paolicchi, E.; Gemignani, F.; Krstic-Demonacos, M.; Dedhar, S.; Mutti, L.; Landi, S. Targeting hypoxic response for cancer therapy. Oncotarget 2016, 7, 13464. [Google Scholar] [CrossRef]
  228. Yu, T.; Tang, B.; Sun, X. Development of inhibitors targeting hypoxia-inducible factor 1 and 2 for cancer therapy. Yonsei Med. J. 2017, 58, 489–496. [Google Scholar] [CrossRef]
  229. Duffy, A.; Melillo, G.; Turkbey, B.; Allen, D.; Choyke, P.; Chen, C.; Raffeld, M.; Doroshow, J.; Murgo, A.; Kummar, S. A pilot trial of oral topotecan (TPT) in patients with refractory advanced solid neoplasms expressing HIF-1α. J. Clin. Oncol. 2010, 28, e13518. [Google Scholar] [CrossRef]
  230. Sounni, N.E.; Noel, A. Targeting the tumor microenvironment for cancer therapy. Clin. Chem. 2013, 59, 85–93. [Google Scholar] [CrossRef]
  231. Fukumura, D.; Jain, R.K. Tumor microenvironment abnormalities: Causes, consequences, and strategies to normalize. J. Cell. Biochem. 2007, 101, 937–949. [Google Scholar] [CrossRef]
  232. Netea-Maier, R.T.; Smit, J.W.; Netea, M.G. Metabolic changes in tumor cells and tumor-associated macrophages: A mutual relationship. Cancer Lett. 2018, 413, 102–109. [Google Scholar] [CrossRef] [PubMed]
  233. Waldmann, T.A. Cytokines in cancer immunotherapy. Cold Spring Harb. Perspect. Biol. 2018, 10, a028472. [Google Scholar] [CrossRef] [PubMed]
  234. Bastos, N.; Ruivo, C.F.; da Silva, S.; Melo, S.A. Exosomes in cancer: Use them or target them? Semin. Cell Dev. Biol. 2018, 78, 13–21. [Google Scholar] [CrossRef] [PubMed]
  235. Luberto, C.; Hassler, D.F.; Signorelli, P.; Okamoto, Y.; Sawai, H.; Boros, E.; Hazen-Martin, D.J.; Obeid, L.M.; Hannun, Y.A.; Smith, G.K. Inhibition of tumor necrosis factor-induced cell death in MCF7 by a novel inhibitor of neutral sphingomyelinase. J. Biol. Chem. 2002, 277, 41128–41139. [Google Scholar] [CrossRef]
  236. Colombo, M.; Raposo, G.; Thery, C. Biogenesis, secretion, and intercellular interactions of exosomes and other extracellular vesicles. Annu. Rev. Cell Dev. Biol. 2014, 30, 255–289. [Google Scholar] [CrossRef]
  237. Datta, A.; Kim, H.; Lal, M.; McGee, L.; Johnson, A.; Moustafa, A.A.; Jones, J.C.; Mondal, D.; Ferrer, M.; Abdel-Mageed, A.B. Manumycin A suppresses exosome biogenesis and secretion via targeted inhibition of Ras/Raf/ERK1/2 signaling and hnRNP H1 in castration-resistant prostate cancer cells. Cancer Lett. 2017, 408, 73–81. [Google Scholar] [CrossRef]
  238. Escrevente, C.; Keller, S.; Altevogt, P.; Costa, J. Interaction and uptake of exosomes by ovarian cancer cells. BMC Cancer 2011, 11, 108. [Google Scholar] [CrossRef]
Figure 1. Birds in a nest: Role of exosomes in intra-prostate cancer (PCa) communications. PCa cells communicate with each other by transfer of exosomal cargo proteins and nucleic acids within the tumor microenvironment. In heterogenic tumor cells and in premetastatic niche, shedding of exosomes from vicious PCa cells reactivate dormant PCa cells to gain new aggressive traits through inducing of cell growth, differentiation, epithelial-mesenchymal transition (EMT), metabolic adaptation under hypoxic conditions, and angiogenesis.
Figure 1. Birds in a nest: Role of exosomes in intra-prostate cancer (PCa) communications. PCa cells communicate with each other by transfer of exosomal cargo proteins and nucleic acids within the tumor microenvironment. In heterogenic tumor cells and in premetastatic niche, shedding of exosomes from vicious PCa cells reactivate dormant PCa cells to gain new aggressive traits through inducing of cell growth, differentiation, epithelial-mesenchymal transition (EMT), metabolic adaptation under hypoxic conditions, and angiogenesis.
Cells 09 00564 g001
Figure 2. The potential roles of prostate cancer (PCa)-associated exosomes in modulating different cells in the TME. Exosomes-associated cargo shuttles bioactive molecules to and from PCa cells to activate stromal cells in the TME for gaining new genetic traits and promoting PCa progression and metastasis. Fibroblast cells (FB), cancer associate fibroblast (CAF), endothelial cells (ECs), patient adipose derived stem cells (pASC), T cell, natural killer cells (NK), lymphocytes, bone marrow-mesenchymal stem cells (BM-MSC), osteoclast cells, extracellular matrix (ECM). Part of this figure was prepared by the aid of Mind the GRAPH program available on
Figure 2. The potential roles of prostate cancer (PCa)-associated exosomes in modulating different cells in the TME. Exosomes-associated cargo shuttles bioactive molecules to and from PCa cells to activate stromal cells in the TME for gaining new genetic traits and promoting PCa progression and metastasis. Fibroblast cells (FB), cancer associate fibroblast (CAF), endothelial cells (ECs), patient adipose derived stem cells (pASC), T cell, natural killer cells (NK), lymphocytes, bone marrow-mesenchymal stem cells (BM-MSC), osteoclast cells, extracellular matrix (ECM). Part of this figure was prepared by the aid of Mind the GRAPH program available on
Cells 09 00564 g002
Figure 3. Effect of stromal cell-derived exosomes on PCa progression and metastasis. Stromal cells release exosomes fully loaded with cargo macromolecules to promote cell proliferation, hypoxic and adaptive metabolic pathways, angiogenesis and metastasis of PCa cells. Cancer-associate fibroblasts (CAFs), endothelial cells (ECs), patient adipose-derived stem cells (pASCs), bone marrow-mesenchymal stem cells (BM-MSC) and osteoclast cells. Part of this figure was prepared by the aid of Mind the GRAPH program.
Figure 3. Effect of stromal cell-derived exosomes on PCa progression and metastasis. Stromal cells release exosomes fully loaded with cargo macromolecules to promote cell proliferation, hypoxic and adaptive metabolic pathways, angiogenesis and metastasis of PCa cells. Cancer-associate fibroblasts (CAFs), endothelial cells (ECs), patient adipose-derived stem cells (pASCs), bone marrow-mesenchymal stem cells (BM-MSC) and osteoclast cells. Part of this figure was prepared by the aid of Mind the GRAPH program.
Cells 09 00564 g003
Table 1. Exosomes-associated cargo proteins and their functional relevance in Tumor microenvironment (TME).
Table 1. Exosomes-associated cargo proteins and their functional relevance in Tumor microenvironment (TME).
Protein(s)Biological FunctionReference(s)
AHNAK Cancer-associated exosomes increase motility of fibroblasts[149]
ANXA2, CLSTN1, FASN, FLNC, FOLH1, and GDF15Correlated with PCa malignancy[67]
CD9Increase the proliferation and chemo-resistance of PCa cells[87,150]
B7-H3Immune checkpoint regulator[151]
CD63, CD81, HSP90, HSP70, TNF1α, IL-6, MMP2, MMP9, Annexin II, TSG101, Akt, ILK1, and β-cateninIncrease stemness, metastasis and CAFs formation[91]
c-Src, IGF-1R and FAKPCa development and angiogenesis[139,140,141]
Ets-1Induce osteoclast differentiation[152]
FasL, TGF-β, galectin-9 and HSP72Evade immune responses[145,146]
Galectin-1Promote angiogenesis[153]
Integrin αvβ3Pro-inflammatory effect on stromal cells[93,137]
Integrin β4, vinculin and P-gpAssociated with taxane and docetaxel resistance [154,155]
MMP14Promote PCa cell growth[156]
PD-L1Immune checkpoint regulator[157]
Rab1a, Rab1b and Rab11aNeoplastic transformation of pASCs[130]
TGF-βDifferentiation of fibroblast to CAFs[131,132,133,134,135]
TIMP-1Associated with PCa aggressiveness [158]
Trop-2, vimentin, N-cadherin and Integrin αvβ3Induce PCa cell invasion[92,136,159,160]
uPAVascularization remodeling of PCa microenvironment[142,143]
VEGFA, HGF and MMPAngiogenesis, EMT and tumor growth[136]
Table 2. PCa-associated exosomes transfer nucleic acids in their cargo, which have significant functions in TME.
Table 2. PCa-associated exosomes transfer nucleic acids in their cargo, which have significant functions in TME.
Nucleic Acid(s)Biological FunctionReference(s)
miR-125b, miR130b, miR-155Neoplastic transformation of PCa patients’ adipose stem cells[130]
miR-125bDownregulate AKT1 expression and induce PCa proliferation[187]
miR-100, miR-21 and miR-139PCa growth and metastasis[176]
miR-141, miR-21, miRNA-375Affect osteoclastogenesis and osteoblastogenesis and help PCa cells to overcome androgen deprivation in long distant metastasis[180,181,188,189]
miR-409, miR-379 and miR-154Support PCa carcinogenesis and metastasis[177]
miR-485-3pAssociated with fludarabine resistance in PCa cells[190]
H-ras, K-ras (mRNA)Neoplastic transformation of PC patients’ adipose stem cells.[130]
LincRNA-p21 (lncRNA)Associated with PC malignancy[186]
lncRNA MY (lncRNA)Promote adjacent cell proliferation and migration[185]
Back to TopTop