9.3B: Cell Signaling and Gene Expression - Biology

9.3B: Cell Signaling and Gene Expression - Biology

We are searching data for your request:

Forums and discussions:
Manuals and reference books:
Data from registers:
Wait the end of the search in all databases.
Upon completion, a link will appear to access the found materials.

Gene expression, vital for cells to function properly, is the process of turning on a gene to produce RNA and protein.

Learning Objectives

  • Describe the regulation of gene expression

Key Points

  • Each cell controls when and how its genes are expressed.
  • Malfunctions in the control of gene expression are detrimental to the cell and can lead to the development of many diseases, such as cancer.
  • In prokaryotic cells, the control of gene expression is mostly at the transcriptional level.
  • In eukaryotic cells, the control of gene expression is at the epigenetic, transcriptional, post-transcriptional, translational, and post-translational levels.

Key Terms

  • translation: a process occurring in the ribosome in which a strand of messenger RNA (mRNA) guides assembly of a sequence of amino acids to make a protein
  • gene expression: the transcription and translation of a gene into messenger RNA and, thus, into a protein
  • transcription: the synthesis of RNA under the direction of DNA

Gene Expression

For a cell to function properly, necessary proteins must be synthesized at the proper time. All cells control or regulate the synthesis of proteins from information encoded in their DNA. The process of turning on a gene to produce RNA and protein is called gene expression.

Whether in a simple unicellular organism or a complex multi-cellular organism, each cell controls when and how its genes are expressed. For this to occur, there must be a mechanism to control when a gene is expressed to make RNA and protein; how much of the protein is made; and when it is time to stop making that protein because it is no longer needed. The regulation of gene expression conserves energy and space. It would require a significant amount of energy for an organism to express every gene at all times, so it is more energy efficient to turn on the genes only when they are required. In addition, only expressing a subset of genes in each cell saves space because DNA must be unwound from its tightly-coiled structure to transcribe and translate the DNA. Cells would have to be enormous if every protein were expressed in every cell all the time. The control of gene expression is extremely complex. Malfunctions in this process are detrimental to the cell and can lead to the development of many diseases, including cancer.

Prokaryotic versus Eukaryotic Gene Expression

To understand how gene expression is regulated, we must first understand how a gene codes for a functional protein in a cell. The process occurs in both prokaryotic and eukaryotic cells, just in slightly different manners. Prokaryotic organisms are single-celled organisms that lack a cell nucleus; their DNA floats freely in the cell cytoplasm. To synthesize a protein, the processes of transcription and translation occur almost simultaneously. When the resulting protein is no longer needed, transcription stops. As a result, the primary method to control what type of protein and how much of each protein is expressed in a prokaryotic cell is the regulation of DNA transcription. All of the subsequent steps occur automatically. When more protein is required, more transcription occurs. Therefore, in prokaryotic cells, the control of gene expression is mostly at the transcriptional level.

Eukaryotic cells, in contrast, have intracellular organelles that add to their complexity. In eukaryotic cells, the DNA is contained inside the cell’s nucleus where it is transcribed into RNA. The newly-synthesized RNA is then transported out of the nucleus into the cytoplasm where ribosomes translate the RNA into protein. The processes of transcription and translation are physically separated by the nuclear membrane: transcription occurs only within the nucleus, and translation occurs only outside the nucleus in the cytoplasm. The regulation of gene expression can occur at all stages of the process. Regulation may occur when the DNA is uncoiled and loosened from nucleosomes to bind transcription factors (epigenetic level); when the RNA is transcribed (transcriptional level); when the RNA is processed and exported to the cytoplasm after it is transcribed (post-transcriptional level); when the RNA is translated into protein (translational level); or after the protein has been made (post-translational level).

Use of Gene‐Manipulated Mice in the Study of Erythropoietin Gene Expression

Transcriptional regulation of animal genes has been classified into two major categories: tissue‐specific and stress‐inducible. Erythropoietin (EPO), an erythroid growth factor, plays a central role in the regulation of red blood cell production. In response to hypoxic and/or anemic stresses, Epo gene expression is markedly induced in kidney and liver thus, the Epo gene has been used as a model for elucidating stress‐inducible gene expression in animals. A key transcriptional regulator of the hypoxia response, hypoxia‐inducible transcription factor (HIF), has been identified and cloned through studies on the Epo gene. Recently developed gene‐modified mouse lines have proven to be a powerful means of exploring the regulatory mechanisms as well as the physiological significance of the tissue‐specific and hypoxia‐inducible expression of the Epo gene. In this chapter, several gene‐modified mouse lines related to EPO and the EPO receptor are introduced, with emphasis placed on the examination of in vivo EPO activity, EPO function in nonhematopoietic tissues, EPO‐producing cells in the kidney, and cis‐acting regulatory elements for Epo gene expression. These in vivo studies of the Epo gene have allowed for a deeper understanding of transcriptional regulation operated in a tissue‐specific and stress‐inducible manner.

An example of an autocrine agent is the cytokine interleukin-1 in monocytes. When interleukin-1 is produced in response to external stimuli, it can bind to cell-surface receptors on the same cell that produced it. [ citation needed ]

Another example occurs in activated T cell lymphocytes, i.e., when a T cell is induced to mature by binding to a peptide:MHC complex on a professional antigen-presenting cell and by the B7:CD28 costimulatory signal. Upon activation, "low-affinity" IL-2 receptors are replaced by "high-affinity" IL-2 receptors consisting of α, β, and γ chains. The cell then releases IL-2, which binds to its own new IL-2 receptors, causing self-stimulation and ultimately a monoclonal population of T cells. These T cells can then go on to perform effector functions such as macrophage activation, B cell activation, and cell-mediated cytoxicity. [ citation needed ]

Tumor development is a complex process that requires cell division, growth, and survival. One approach used by tumors to upregulate growth and survival is through autocrine production of growth and survival factors. Autocrine signaling plays critical roles in cancer activation and also in providing self-sustaining growth signals to tumors. [ citation needed ]

In the Wnt pathway Edit

Normally, the Wnt signaling pathway leads to stabilization of β-catenin through inactivation of a protein complex containing the tumor suppressors APC and Axin. This destruction complex normally triggers β-catenin phosphorylation, inducing its degradation. De-regulation of the autocrine Wnt signaling pathway via mutations in APC and Axin have been linked to activation of various types of human cancer. [2] [3] Genetic alterations that lead to de-regulation of the autocrine Wnt pathway result in transactivation of epidermal growth factor receptor (EGFR) and other pathways, in turn contributing to proliferation of tumor cells. In colorectal cancer, for example, mutations in APC, axin, or β-catenin promote β-catenin stabilization and transcription of genes encoding cancer-associated proteins. Furthermore, in human breast cancer, interference with the de-regulated Wnt signaling pathway reduces proliferation and survival of cancer. These findings suggest that interference with Wnt signaling at the ligand-receptor level may improve the effectiveness of cancer therapies. [3]

IL-6 Edit

Interleukin 6 (acronym: IL-6) is a cytokine that is important for many aspects of cellular biology including immune responses, cell survival, apoptosis, as well as proliferation. [4] Several studies have outlined the importance of autocrine IL-6 signaling in lung and breast cancers. For example, one group found a positive correlation between persistently activated tyrosine-phosphorylated STAT3 (pSTAT3), found in 50% of lung adenocarcinomas, and IL-6. Further investigation revealed that mutant EGFR could activate the oncogenic STAT3 pathway via upregulated IL-6 autocrine signaling. [5]

Similarly, HER2 overexpression occurs in approximately a quarter of breast cancers and correlates with poor prognosis. Recent research revealed that IL-6 secretion induced by HER2 overexpression activated STAT3 and altered gene expression, resulting in an autocrine loop of IL-6/STAT3 expression. Both mouse and human in vivo models of HER2-overexpressing breast cancers relied critically on this HER2–IL-6–STAT3 signaling pathway. [6] Another group found that high serum levels of IL-6 correlated with poor outcome in breast cancer tumors. Their research showed that autocrine IL-6 signaling induced malignant features in Notch-3 expressing mammospheres. [7]

IL-7 Edit

A study demonstrates how the autocrine production of the IL-7 cytokine mediated by T-cell acute lymphoblastic leukemia (T-ALL) can be involved in the oncogenic development of T-ALL and offer novel insights into T-ALL spreading. [8]


Another agent involved in autocrine cancer signaling is vascular endothelial growth factor (VEGF). VEGF, produced by carcinoma cells, acts through paracrine signaling on endothelial cells and through autocrine signaling on carcinoma cells. [9] Evidence shows that autocrine VEGF is involved in two major aspects of invasive carcinoma: survival and migration. Moreover, it was shown that tumor progression selects for cells that are VEGF-dependent, challenging the belief that VEGF's role in cancer is limited to angiogenesis. Instead, this research suggests that VEGF receptor-targeted therapeutics may impair cancer survival and invasion as well as angiogenesis. [9] [10]

Promotion of metastasis Edit

Metastasis is a major cause of cancer deaths, and strategies to prevent or halt invasion are lacking. One study showed that autocrine PDGFR signaling plays an essential role in epithelial-mesenchymal transition (EMT) maintenance in vitro, which is known to correlate well with metastasis in vivo. The authors showed that the metastatic potential of oncogenic mammary epithelial cells required an autocrine PDGF/PDGFR signaling loop, and that cooperation of autocrine PDGFR signaling with oncogenic was required for survival during EMT. Autocrine PDGFR signaling also contributes to maintenance of EMT, possibly through activation of STAT1 and other distinct pathways. In addition, expression of PDGFRα and -β correlated with invasive behavior in human mammary carcinomas. [11] This indicates the numerous pathways through which autocrine signaling can regulate metastatic processes in a tumor.

Development of therapeutic targets Edit

The growing knowledge behind the mechanism of autocrine signaling in cancer progression has revealed new approaches for therapeutic treatment. For example, autocrine Wnt signaling could provide a novel target for therapeutic intervention by means of Wnt antagonists or other molecules that interfere with ligand-receptor interactions of the Wnt pathway. [2] [3] In addition, VEGF-A production and VEGFR-2 activation on the surface of breast cancer cells indicates the presence of a distinct autocrine signaling loop that enables breast cancer cells to promote their own growth and survival by phosphorylation and activation of VEGFR-2. This autocrine loop is another example of an attractive therapeutic target. [9]

In HER2 overexpressing breast cancers, the HER2–IL-6–STAT3 signaling relationship could be targeted to develop new therapeutic strategies. [6] HER2 kinase inhibitors, such as lapatinib, have also demonstrated clinical efficacy in HER2 overexpressing breast cancers by disrupting a neuregulin-1 (NRG1)-mediated autocrine loop. [12]

In the case of PDGFR signalling, overexpression of a dominant-negative PDGFR or application of the cancer drug STI571 are both approaches being explored to therapeutically interference with metastasis in mice. [11]

In addition, drugs may be developed that activate autocrine signaling in cancer cells that would not otherwise occur. For example, a small-molecule mimetic of Smac/Diablo that counteracts the inhibition of apoptosis has been shown to enhance apoptosis caused by chemotherapeutic drugs through autocrine-secreted tumor necrosis factor alpha (TNFα). In response to autocrine TNFα signaling, the Smac mimetic promotes formation of a RIPK1-dependent caspase-8-activating complex, leading to apoptosis. [13]

Role in drug resistance Edit

Recent studies have reported the ability of drug-resistant cancer cells to acquire mitogenic signals from previously neglected autocrine loops, causing tumor recurrence.

For example, despite widespread expression of epidermal growth factor receptors (EGFRs) and EGF family ligands in non-small-cell lung cancer (NSCLC), EGFR-specific tyrosine kinase inhibitors such as gefitinib have shown limited therapeutic success. This resistance is proposed to be because autocrine growth signaling pathways distinct from EGFR are active in NSCLC cells. Gene expression profiling revealed the prevalence of specific fibroblast growth factors (FGFs) and FGF receptors in NSCLC cell lines, and found that FGF2, FGF9 and their receptors encompass a growth factor autocrine loop that is active in a subset of gefitinib-resistant NSCLC cell lines. [14]

In breast cancer, the acquisition of tamoxifen resistance is another major therapeutic problem. It has been shown that phosphorylation of STAT3 and RANTES expression are increased in response to tamoxifen in human breast cancer cells. In a recent study, one group showed that STAT3 and RANTES contribute to the maintenance of drug resistance by upregulating anti-apoptotic signals and inhibiting caspase cleavage. These mechanisms of STAT3-RANTES autocrine signaling suggest a novel strategy for management of patients with tamoxifen-resistant tumors. [15]

3B, a novel photosensitizer, inhibits glycolysis and inflammation via miR-155-5p and breaks the JAK/STAT3/SOCS1 feedback loop in human breast cancer cells

Compared to normal cells, most cancer cells produce ATP by glycolysis under aerobic conditions rather than via the tricarboxylic acid cycle (TCA). This study is intended to determine whether 3B, a novel photosensitizer, can inhibit glycolysis and inflammation in breast cancer cells. We showed that 3B had the ability to repress glucose consumption as well as the generation of ATP, lactate and lactate dehydrogenase. 3B-PDT not only inhibited the expression of IL-1β and IL-6 but also affected the JAK-STAT3 inflammatory pathway in vitro. The present study showed that 3B featured a significant inhibitory effect on the expression of microRNA-155-5p and SOCS1 might serve as a target gene. In vivo studies revealed that 3B inhibited tumor growth and exhibited almost no side effects. Therefore, through the anti-glycolytic effect and breakage of the JAK/STAT3/SOCS1 feedback loop via miR-155-5p, 3B may potentially serve as a potential therapeutic agent against breast cancer.

Evolving DNA methylation and gene expression markers of B-cell chronic lymphocytic leukemia are present in pre-diagnostic blood samples more than 10 years prior to diagnosis

Background: B-cell chronic lymphocytic leukemia (CLL) is a common type of adult leukemia. It often follows an indolent course and is preceded by monoclonal B-cell lymphocytosis, an asymptomatic condition, however it is not known what causes subjects with this condition to progress to CLL. Hence the discovery of prediagnostic markers has the potential to improve the identification of subjects likely to develop CLL and may also provide insights into the pathogenesis of the disease of potential clinical relevance.

Results: We employed peripheral blood buffy coats of 347 apparently healthy subjects, of whom 28 were diagnosed with CLL 2.0-15.7 years after enrollment, to derive for the first time genome-wide DNA methylation, as well as gene and miRNA expression, profiles associated with the risk of future disease. After adjustment for white blood cell composition, we identified 722 differentially methylated CpG sites and 15 differentially expressed genes (Bonferroni-corrected p < 0.05) as well as 2 miRNAs (FDR < 0.05) which were associated with the risk of future CLL. The majority of these signals have also been observed in clinical CLL, suggesting the presence in prediagnostic blood of CLL-like cells. Future CLL cases who, at enrollment, had a relatively low B-cell fraction (<10%), and were therefore less likely to have been suffering from undiagnosed CLL or a precursor condition, showed profiles involving smaller numbers of the same differential signals with intensities, after adjusting for B-cell content, generally smaller than those observed in the full set of cases. A similar picture was obtained when the differential profiles of cases with time-to-diagnosis above the overall median period of 7.4 years were compared with those with shorted time-to-disease. Differentially methylated genes of major functional significance include numerous genes that encode for transcription factors, especially members of the homeobox family, while differentially expressed genes include, among others, multiple genes related to WNT signaling as well as the miRNAs miR-150-5p and miR-155-5p.

Conclusions: Our findings demonstrate the presence in prediagnostic blood of future CLL patients, more than 10 years before diagnosis, of CLL-like cells which evolve as preclinical disease progresses, and point to early molecular alterations with a pathogenetic potential.

Keywords: Biomarkers of risk Epigenomics Molecular epidemiology Prospective cohort Transcriptomics miRNA.

Conflict of interest statement

Ethics approval and consent to participate

The EnviroGenomarkers project and its associated studies and experimental protocols were approved by the Regional Ethical Review Board of the Umeå Division of Medical Research, for the Swedish cohort, and the Florence Health Unit Local Ethical Committee, for the Italian cohort. All participants gave written informed consent.

Consent for publication
Competing interests

The authors declare that they have no competing interests.

Publisher’s Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.


Carboxy terminal truncation of the G-CSF-R, as seen in patients with AML/SCN, leads to augmented and prolonged activation of Stat5 by G-CSF, suggesting that the carboxy terminus of the G-CSF-R negatively regulates Stat5 activation. It has been shown that Shp-1 reduces the intensity but not the duration of G-CSF-stimulated Stat5 activation, and the carboxy terminal region of the G-CSF-R but not the tyrosine residues located in this region is required for the inhibitory action of Shp-1 [ 21 ]. In this paper, we have investigated the mechanism by which the carboxy terminus of the G-CSF-R regulates the duration of Stat5 activation. Here, we provide the first evidence that Tyr 729 of the G-CSF-R controls the duration but not the intensity of G-CSF-stimulated Stat5 activation. We also demonstrate that SOCS1 and SOCS3 attenuate G-CSF-R signaling, and Tyr 729 of the G-CSF-R is required for the maximal inhibitory effects of SOCS1 and SOCS3.

Our results are consistent with recent studies showing that SOCS3 binds to Tyr 729 of the G-CSF-R and inhibits Stat-dependent luciferase activity stimulated by G-CSF [ 35 36 37 ]. However, these studies did not examine Stat5 tyrosine phosphorylation and DNA binding activity in hematopoietic cells, and therefore, it remains unknown as to which aspects of Stat5 activation are affected by SOCS3. The data presented here strongly suggest that SOCS3 and SOCS1 control the duration, but not the magnitude, of Stat5 activation in hematopoietic cells. It appears that Tyr 729 controls the functions of SOCS3 and SOCS1 by at least two distinct mechanisms. Removal of Tyr 729 from the G-CSF-R through carboxy terminal truncation or amino acid substitution not only delays the induction of SOCS3 and SOCS1 transcripts by G-CSF but also diminishes the negative effect of SOCS3 and SOCS1 on Stat5 activation. Thus, Tyr 729 of the G-CSF-R regulates the rates at which SOCS3 and SOCS1 are up-regulated by G-CSF and the efficacies with which the two SOCS proteins inhibit G-CSF-R signaling. Conceivably, mutation of Tyr 729 causes prolonged G-CSF-R signaling by disrupting both negative regulatory mechanisms.

In addition to deregulated Stat5 activation, carboxy terminal truncation of the G-CSF-R results in sustained activation of Akt by G-CSF [ 38 , 39 ]. Our results indicate that Tyr 729 of the G-CSF-R is critically involved in controlling the duration of G-CSF-stimulated activation of Akt and Erk1/2. Activation of Akt and Erk1/2 by G-CSF was also prolonged upon inhibition of protein or RNA synthesis, suggesting that SOCS1 and SOCS3 may also exert a negative effect on the activation of Akt and Erk1/2. Indeed, expression of SOCS1 or SOCS3 in 293T cells attenuated G-CSF-induced Akt phosphorylation (data not shown). It is likely that SOCS1 and SOCS3 may target a common upstream component of the Stat5, Akt, and Erk1/2 pathways, although one cannot exclude the possibility that the two SOCS proteins may directly inhibit the activation of individual components within the three pathways. As the Jak and Src family kinases are differentially required for G-CSF-stimulated activation of Stat5 and Akt, respectively [ 38 ], one potential, common target of SOCS1 and SOCS3 is the G-CSF-R. In support of this, we have observed that tyrosine phosphorylation of the G-CSF-R induced by G-CSF was completely blocked upon expression of SOCS1 or SOCS3 in 293T cells (data not shown).

A recent study showed that CIS was also up-regulated by G-CSF and bound to phosphopeptides corresponding to Tyr 729 and Tyr 744 of the G-CSFR [ 40 ]. However, we consistently observed that expression of CIS enhanced Stat5 activation by G-CSF in 293T cells, consistent with the report by van de Geijn et al. [ 37 ]. It remains to be determined, with respect to the relative contribution of SOCS1 and SOCS3 to the attenuation of G-CSF-R signaling and the precise mechanism by which the two SOCS proteins down-regulate the activation of G-CSF-stimulated pathways. In addition to Tyr 729, SOCS3 has been shown to bind to Tyr 704 of the G-CSF-R with a reduced affinity [ 35 ], which may explain the partial inhibitory effect of SOCS3 on Stat5 activation mediated by the Y729F mutant. In contrast to SOCS3, it is generally believed that SOCS1 inhibits cytokine receptor signaling via interacting directly with the Jak family kinases but not with the cytokine receptors [ 34 , 41 ]. In fact, it has been shown that SOCS1 fails to bind to any of the cytoplasmic tyrosine residues including Tyr 729 of the G-CSF-R [ 35 ]. We also have been unable to detect any interaction between SOCS1 and the G-CSF-R (unpublished data). However, our data clearly show that Tyr 729 of the G-CSF-R is required for the maximal inhibitory effect of SOCS1 on G-CSF-stimulated Stat5 activation. Whether SOCS1 may interact indirectly with Tyr 729 of the G-CSF-R remains to be examined.

Regardless of the mechanisms whereby SOCS3 and SOCS1 exert their negative effect on G-CSF-R signaling, the results presented here together with the previously report [ 21 ] demonstrate that the intensity and duration of G-CSF-stimulated Stat5 activation are regulated by at least two distinct regulatory mechanisms, i.e., the one mediated by Shp-1, which reduces the magnitude of Stat5 activation, and the one mediated by SOCS3 and/or SOCS1, which controls the duration of Stat5 activation (Fig. 9). To our knowledge, such a regulatory feature has not been reported for other cytokine receptors. Additionally, our data reveal an important molecular mechanism that explains, at least in part, why truncation of the carboxy terminal region of the G-CSF-R leads to prolonged receptor signaling and may thus contribute to leukemogenesis.

Expression of the different G-CSF-R in 32D cells. (A) Schematic representation of the different forms of the G-CSF-R. Shown are the transmembrane and cytoplasmic domains. Boxes 1–3 denote regions that are conserved among some members of the cytokine receptor superfamily. Cytoplasmic tyrosine residues are also indicated. (B) Flow cytometric analysis of G-CSF-R expression in 32D cells. Cells were incubated sequentially with the anti G-CSF-R antibody and the fluorophore-labeled goat anti-mouse IgG antibody at 4°C. Samples were analyzed by flow cytometry. Ctr, control.

Activation of Stat5, Akt, and Erk1/2 by G-CSF in 32D cells expressing the different forms of the G-CSF-R. (A) 32D cells expressing the WT, D715, or mA receptor were starved in serum-free medium for 4 h prior to G-CSF stimulation for 15 min. Phosphorylation (p) of Stat5, Akt, and Erk1/2 was determined by Western blotting using the phospho-specific antibodies. The membranes were reprobed with antibodies to Stat5, Akt, and Erk1/2. (B) Survival of 32D cells upon G-CSF withdrawal. Cells, as indicated, were incubated with G-CSF for 8 h and then cultured in quadruplicate in medium containing no growth factors. Cell viability was determined by exclusion of trypan blue staining at the indicated times. The data are presented as mean ± sd of quadruplicate determinations. ∗, Significant differences as compared with the survival of 32D/WT cells (Student’s t-test P<0.05). Comparable results were obtained in four independent experiments.

Activation of Stat5, Akt, and Erk1/2 by the different G-CSF-R forms. 32D and Ba/F3 cells expressing the different forms of the G-CSF-R were stimulated with G-CSF after starvation in serum-free medium for 4 h. (A) Activation of Stat5, Akt, and Erk1/2 in 32D cells was examined by immunoblotting with the phospho-specific antibodies as indicated. The membranes were subsequently probed with antibodies to Stat5, Akt, and Erk1/2. (B) Stat5 DNA binding activity in 32D cells was assessed by EMSA using the β-casein probe. (C) Activation of Stat5 in Ba/F3 cells was determined by immunoblotting with the phospho-specific Stat5 antibody. The membrane was reprobed for Stat5.

Effects of actinomycin D and cycloheximide on G-CSF-induced activation of Stat5, Akt, and Erk1/2. After starvation in serum-free medium for 2 h, 32D/WT cells were treated with actinomycin D (5 μg/ml) or cycloheximide (30 μg/ml) for an additional 2 h prior to stimulation with G-CSF. Whole cell extracts were prepared and examined for Stat5 tyrosine phosphorylation and DNA binding activity using the β-casein probe and for Akt and Erk1/2 phosphorylation. The membranes were reprobed with the antibodies to Stat5, Akt, and Erk1/2.

Effects of the different SOCS proteins on G-CSF-induced Stat5 activation. 293T cells were transiently transfected with cDNAs encoding the WT G-CSF-R and Stat5a, together with cDNAs encoding the different FLAG-tagged SOCS proteins or the empty vector. Twenty hours after transfection, cells were starved for 4 h and stimulated with G-CSF for the indicated times. Whole cell extracts were examined by Western blotting (WB) for Stat5 tyrosine phosphorylation (p-Stat5 top panel) and for expression of Stat5 (middle panel) and the different SOCS proteins using the anti-FLAG antibody (bottom panel).

Northern blot analysis of SOCS1 and SOCS3 transcripts. (A) 32D cells expressing the WT, D715, and Y729F forms of the G-CSF-R were stimulated with G-CSF for the indicated times following starvation in serum-free medium for 4 h. Total RNA extracted from the cells was electrophoresed in a 1% agarose gel, blotted to membrane, and probed with [γ- 32 P]-labeled probes for SOCS1 (top panel) and SOCS3 (middle panel). Sample loadings were determined by ethidium bromide staining of 18S rRNA (bottom panel). (B and C) The intensities of the bands for SOCS1 and SOCS3 transcripts were quantitated using a phosphoimager.

Requirement of Tyr 729 of the G-CSF-R for SOCS1- and SOCS3-mediated inhibition of Stat5 activation. 293T cells were transiently transfected with Stat5a expression construct and the expression constructs for the WT and the different tyrosine-to-phenylalanine substitution forms of the G-CSF-R together with constructs for SOCS1, SOCS3, or empty vector. G-CSF-induced tyrosine phosphorylation of Stat5 was examined by Western blotting using the phospho-specific anti-Stat5 antibody. Expression of SOCS1 and SOCS3 was determined using the anti-FLAG antibody.

Effects of SOCS1 and SOCS3 on G-CSF-dependent proliferation. BaF/WT and BaF/Y729F cells were transiently transfected with the empty vector (Ctr) or expression constructs for SOCS1 and SOCS3. Six hours after transfection, cells were washed and incubated in triplicate in serum-free medium for 24 h in the presence of G-CSF (20 ng/ml). Cells were then pulsed with MTS for 2 h, and MTS uptake was determined. Data are presented as percentage (mean± sd ) of MTS uptake by cells transfected with the empty vector. Comparable results were obtained in three independent experiments. The differences in the rate of inhibition by SOCS1 and SOCS3 are statistically significant between BaF/WT and BaF/Y729F cells (Student’s t-test P<0.01).

A model of negative regulation of G-CSF-stimulated Stat5 activation. The G-CSF-R form dimers or oligomers upon ligand binding. Shp-1 is constitutively expressed in myeloid cells and controls the magnitude of Stat5 activation via interacting with the carboxy terminus of the G-CSF-R. Carboxy terminal tyrosine residues of the receptor appear dispensable for Shp-1 action [ 21 , 22 ]. SOCS3 and SOCS1 are up-regulated rapidly by signals transduced from the G-CSF-R and control the duration of Stat5 activation. Tyr 729 of the G-CSF-R is required for the maximal inhibitory activities of SOCS3 and SOCS1.


There are four JAK proteins: JAK1, JAK2, JAK3 and TYK2. [1] JAKs contains a FERM domain (approximately 400 residues), an SH2-related domain (approximately 100 residues), a kinase domain (approximately 250 residues) and a pseudokinase domain (approximately 300 residues). [2] The kinase domain is vital for JAK activity, since it allows JAKs to phosphorylate (add phosphate groups to) proteins.

There are seven STAT proteins: STAT1, STAT2, STAT3, STAT4, STAT5A, STAT5B and STAT6. [1] STAT proteins contain many different domains, each with a different function, of which the most conserved region is the SH2 domain. [2] The SH2 domain is formed of 2 α-helices and a β-sheet and is formed approximately from residues 575–680. [2] [3] STATs also have transcriptional activation domains (TAD), which are less conserved and are located at the C-terminus. [4] In addition, STATs also contain: tyrosine activation, amino-terminal, linker, coiled-coil and DNA-binding domains. [4]

The binding of various ligands, usually cytokines, such as interferons and interleukins, to cell-surface receptors, causes the receptors to dimerize, which brings the receptor-associated JAKs into close proximity. [6] The JAKs then phosphorylate each other on tyrosine residues located in regions called activation loops, through a process called transphosphorylation, which increases the activity of their kinase domains. [6] The activated JAKs then phosphorylate tyrosine residues on the receptor, creating binding sites for proteins possessing SH2 domains. [6] STATs then bind to the phosphorylated tyrosines on the receptor using their SH2 domains, and then they are tyrosine-phosphorylated by JAKs, causing the STATs to dissociate from the receptor. [2] At least STAT5 requires glycosylation at threonine 92 for strong STAT5 tyrosine phosphorylation. [7] These activated STATs form hetero- or homodimers, where the SH2 domain of each STAT binds the phosphorylated tyrosine of the opposite STAT, and the dimer then translocates to the cell nucleus to induce transcription of target genes. [2] STATs may also be tyrosine-phosphorylated directly by receptor tyrosine kinases - but since most receptors lack built-in kinase activity, JAKs are usually required for signalling. [1]

Movement of STATs from the cytosol to the nucleus Edit

To move from the cytosol to the nucleus, STAT dimers have to pass through nuclear pore complexes (NPCs), which are protein complexes present along the nuclear envelope that control the flow of substances in and out of the nucleus. To enable STATs to move into the nucleus, an amino acid sequence on STATs, called the nuclear localization signal (NLS), is bound by proteins called importins. [4] Once the STAT dimer (bound to importins) enters the nucleus, a protein called Ran (associated with GTP) binds to the importins, releasing them from the STAT dimer. [8] The STAT dimer is then free in the nucleus.

Specific STATs appear to bind to specific importin proteins. For example, STAT3 proteins can enter the nucleus by binding to importin α3 and importin α6. [9] On the other hand, STAT1 and STAT2 bind to importin α5. [4] Studies indicate that STAT2 requires a protein called interferon regulatory factor 9 (IRF9) to enter the nucleus. [8] Not as much is known about nuclear entrance of other STATs, but it has been suggested that a sequence of amino acids in the DNA-binding domain of STAT4 might allow nuclear import also, STAT5 and STAT6 can both bind to importin α3. [8] In addition, STAT3, STAT5 and STAT6 can enter the nucleus even if they are not phosphorylated at tyrosine residues. [8]

Role of post-translational modifications Edit

After STATs are made by protein biosynthesis, they have non-protein molecules attached to them, called post-translational modifications. One example of this is tyrosine phosphorylation (which is fundamental for JAK-STAT signalling), but STATs experience other modifications, which may affect STAT behaviour in JAK-STAT signalling. These modifications include: methylation, acetylation and serine phosphorylation.

  • Methylation. STAT3 can be dimethylated (have two methyl groups) on a lysine residue, at position 140, and it is suggested that this could reduce STAT3 activity. [10] There is debate as to whether STAT1 is methylated on an arginine residue (at position 31), and what the function of this methylation could be. [11]
  • Acetylation. STAT1, STAT2, STAT3, STAT5 and STAT6 have been shown to be acetylated. [12] STAT1 may have an acetyl group attached to lysines at positions 410 and 413, and as a result, STAT1 can promote the transcription of apoptotic genes - triggering cell death. [12] STAT2 acetylation is important for interactions with other STATs, and for the transcription of anti-viral genes. [4]

Acetylation of STAT3 has been suggested to be important for its dimerization, DNA-binding and gene-transcribing ability, and IL-6 JAK-STAT pathways that use STAT3 require acetylation for transcription of IL-6 response genes. [12] STAT5 acetylation on lysines at positions 694 and 701 is important for effective STAT dimerization in prolactin signalling. [13] Adding acetyl groups to STAT6 is suggested to be essential for gene transcription in some forms of IL-4 signalling, but not all the amino acids which are acetylated on STAT6 are known. [12]

  • Serine phosphorylation. Most of the seven STATs (except STAT2) undergo serine phosphorylation. [2] Serine phosphorylation of STATs has been shown to reduce gene transcription. [14] It is also required for the transcription of some target genes of the cytokines IL-6 and IFN- γ. [11] It has been proposed that phosphorylation of serine can regulate STAT1 dimerization, [11] and that continuous serine phosphorylation on STAT3 influences cell division. [15]

Recruitment of co-activators Edit

Like many other transcription factors, STATs are capable of recruiting co-activators such as CBP and p300, and these co-activators increase the rate of transcription of target genes. [2] The coactivators are able to do this by making genes on DNA more accessible to STATs and by recruiting proteins needed for transcription of genes. The interaction between STATs and coactivators occurs through the transactivation domains (TADs) of STATs. [2] The TADs on STATs can also interact with histone acetyltransferases (HATs) [16] these HATs add acetyl groups to lysine residues on proteins associated with DNA called histones. Adding acetyl groups removes the positive charge on lysine residues, and as a result there are weaker interactions between histones and DNA, making DNA more accessible to STATs and enabling an increase in the transcription of target genes.

Integration with other signalling pathways Edit

JAK-STAT signalling is able to interconnect with other cell-signalling pathways, such as the PI3K/AKT/mTOR pathway. [17] When JAKs are activated and phosphorylate tyrosine residues on receptors, proteins with SH2 domains (such as STATs) are able bind to the phosphotyrosines, and the proteins can carry out their function. Like STATs, the PI3K protein also has an SH2 domain, and therefore it is also able to bind to these phosphorylated receptors. [17] As a result, activating the JAK-STAT pathway can also activate PI3K/AKT/mTOR signalling.

JAK-STAT signalling can also integrate with the MAPK/ERK pathway. Firstly, a protein important for MAPK/ERK signalling, called Grb2, has an SH2 domain, and therefore it can bind to receptors phosphorylated by JAKs (in a similar way to PI3K). [17] Grb2 then functions to allow the MAPK/ERK pathway to progress. Secondly, a protein activated by the MAPK/ERK pathway, called MAPK (mitogen-activated protein kinase), can phosphorylate STATs, which can increase gene transcription by STATs. [17] However, although MAPK can increase transcription induced by STATs, one study indicates that phosphorylation of STAT3 by MAPK can reduce STAT3 activity. [18]

One example of JAK-STAT signalling integrating with other pathways is Interleukin-2 (IL-2) receptor signaling in T cells. IL-2 receptors have γ (gamma) chains, which are associated with JAK3, which then phosphorylates key tyrosines on the tail of the receptor. [19] Phosphorylation then recruits an adaptor protein called Shc, which activates the MAPK/ERK pathway, and this facilitates gene regulation by STAT5. [19]

Alternative signalling pathway Edit

An alternative mechanism for JAK-STAT signalling has also been suggested. In this model, SH2 domain-containing kinases, can bind to phosphorylated tyrosines on receptors and directly phosphorylate STATs, resulting in STAT dimerization. [6] Therefore, unlike the traditional mechanism, STATs can be phosphorylated not just by JAKs, but by other receptor-bound kinases. So, if one of the kinases (either JAK or the alternative SH2-containing kinase) cannot function, signalling may still occur through activity of the other kinase. [6] This has been shown experimentally. [20]

Given that many JAKs are associated with cytokine receptors, the JAK-STAT signalling pathway plays a major role in cytokine receptor signalling. Since cytokines are substances produced by immune cells that can alter the activity of neighbouring cells, the effects of JAK-STAT signalling are often more highly seen in cells of the immune system. For example, JAK3 activation in response to IL-2 is vital for lymphocyte development and function. [21] Also, one study indicates that JAK1 is needed to carry out signalling for receptors of the cytokines IFNγ, IL-2, IL-4 and IL-10. [22]

The JAK-STAT pathway in cytokine receptor signalling can activate STATs, which can bind to DNA and allow the transcription of genes involved in immune cell division, survival, activation and recruitment. For example, STAT1 can enable the transcription of genes which inhibit cell division and stimulate inflammation. [2] Also, STAT4 is able to activate NK cells (natural killer cells), and STAT5 can drive the formation of white blood cells. [2] [23] In response to cytokines, such as IL-4, JAK-STAT signalling is also able to stimulate STAT6, which can promote B-cell proliferation, immune cell survival, and the production of an antibody called IgE. [2]

JAK-STAT signalling plays an important role in animal development. The pathway can promote blood cell division, as well as differentiation (the process of a cell becoming more specialised). [24] In some flies with faulty JAK genes, too much blood cell division can occur, potentially resulting in leukaemia. [25] JAK-STAT signalling has also been associated with excessive white blood cell division in humans and mice. [24]

The signalling pathway is also crucial for eye development in the fruit fly (Drosophila melanogaster). When mutations occur in genes coding for JAKs, some cells in the eye may be unable to divide, and other cells, such as photoreceptor cells, have been shown not to develop correctly. [24]

The entire removal of a JAK and a STAT in Drosophila causes death of Drosophila embryos, whilst mutations in the genes coding for JAKs and STATs can cause deformities in the body patterns of flies, particularly defects in forming body segments. [24] One theory as to how interfering with JAK-STAT signalling might cause these defects is that STATs may directly bind to DNA and promote the transcription of genes involved in forming body segments, and therefore by mutating JAKs or STATs, flies experience segmentation defects. [26] STAT binding sites have been identified on one of these genes, called even-skipped (eve), to support this theory. [27] Of all the segment stripes affected by JAK or STAT mutations, the fifth stripe is affected the most, the exact molecular reasons behind this are still unknown. [24]

Given the importance of the JAK-STAT signalling pathway, particularly in cytokine signalling, there are a variety of mechanisms that cells possess to regulate the amount of signalling that occurs. Three major groups of proteins that cells use to regulate this signalling pathway are protein inhibitors of activated STAT (PIAS), [28] protein tyrosine phosphatases (PTPs) [29] and suppressors of cytokine signalling (SOCS). [30]

Protein inhibitors of activated STATs (PIAS) Edit

PIAS are a four-member protein family made of: PIAS1, PIAS3, PIASx, and PIASγ. [31] The proteins add a marker, called SUMO (small ubiquitin-like modifier), onto other proteins – such as JAKs and STATs, modifying their function. [31] The addition of a SUMO group onto STAT1 by PIAS1 has been shown to prevent activation of genes by STAT1. [32] Other studies have demonstrated that adding a SUMO group to STATs may block phosphorylation of tyrosines on STATs, preventing their dimerization and inhibiting JAK-STAT signalling. [33] PIASγ has also been shown to prevent STAT1 from functioning. [34] PIAS proteins may also function by preventing STATs from binding to DNA (and therefore preventing gene activation), and by recruiting proteins called histone deacetylases (HDACs), which lower the level of gene expression. [31]

Protein tyrosine phosphatases (PTPs) Edit

Since adding phosphate groups on tyrosines is such an important part of how the JAK-STAT signalling pathway functions, removing these phosphate groups can inhibit signalling. PTPs are tyrosine phosphatases, so are able to remove these phosphates and prevent signalling. Three major PTPs are SHP-1, SHP-2 and CD45. [35]

    . SHP-1 is mainly expressed in blood cells. [36] It contains two SH2 domains and a catalytic domain (the region of a protein that carries out the main function of the protein) - the catalytic domain contains the amino acid sequence VHCSAGIGRTG (a sequence typical of PTPs). [37] As with all PTPs, a number of amino acid structures are essential for their function: conserved cysteine, arginine and glutamine amino acids, and a loop made of tryptophan, proline and aspartate amino acids (WPD loop). [37] When SHP-1 is inactive, the SH2 domains interact with the catalytic domain, and so the phosphatase is unable to function. [37] When SHP-1 is activated however, the SH2 domains move away from the catalytic domain, exposing the catalytic site and therefore allowing phosphatase activity. [37] SHP-1 is then able to bind and remove phosphate groups from the JAKs associated with receptors, preventing the transphosphorylation needed for the signalling pathway to progress.

One example of this is seen in the JAK-STAT signalling pathway mediated by the erythropoietin receptor (EpoR). Here, SHP-1 binds directly to a tyrosine residue (at position 429) on EpoR and removes phosphate groups from the receptor-associated JAK2. [38] The ability of SHP-1 to negatively regulate the JAK-STAT pathway has also been seen in experiments using mice lacking SHP-1. [39] These mice experience characteristics of autoimmune diseases and show high levels of cell proliferation, which are typical characteristics of an abnormally high level of JAK-STAT signalling. [39] Additionally, adding methyl groups to the SHP-1 gene (which reduces the amount of SHP-1 produced) has been linked to lymphoma (a type of blood cancer) . [40]

However, SHP-1 may also promote JAK-STAT signalling. A study in 1997 found that SHP-1 potentially allows higher amounts of STAT activation, as opposed to reducing STAT activity. [41] A detailed molecular understanding for how SHP-1 can both activate and inhibit the signalling pathway is still unknown. [35]

    . SHP-2 has a very similar structure to SHP-1, but unlike SHP-1, SHP-2 is produced in many different cell types - not just blood cells. [42] Humans have two SHP-2 proteins, each made up of 593 and 597 amino acids. [37] The SH2 domains of SHP-2 appear to play an important role in controlling the activity of SHP-2. One of the SH2 domains binds to the catalytic domain of SHP-2, to prevent SHP-2 functioning. [35] Then, when a protein with a phosphorylated tyrosine binds, the SH2 domain changes orientation and SHP-2 is activated. [35] SHP-2 is then able to remove phosphate groups from JAKs, STATs and the receptors themselves - so, like SHP-1, can prevent the phosphorylation needed for the pathway to continue, and therefore inhibit JAK-STAT signalling. Like SHP-1, SHP-2 is able to remove these phosphate groups through the action of the conserved cysteine, arginine, glutamine and WPD loop. [37]

Negative regulation by SHP-2 has been reported in a number of experiments - one example has been when exploring JAK1/STAT1 signalling, where SHP-2 is able to remove phosphate groups from proteins in the pathway, such as STAT1. [43] In a similar manner, SHP-2 has also been shown to reduce signalling involving STAT3 and STAT5 proteins, by removing phosphate groups. [44] [45]

Like SHP-1, SHP-2 is also believed to promote JAK-STAT signalling in some instances, as well as inhibit signalling. For example, one study indicates that SHP-2 may promote STAT5 activity instead of reducing it. [46] Also, other studies propose that SHP-2 may increase JAK2 activity, and promote JAK2/STAT5 signalling. [47] It is still unknown how SHP2 can both inhibit and promote JAK-STAT signalling in the JAK2/STAT5 pathway one theory is that SHP-2 may promote activation of JAK2, but inhibit STAT5 by removing phosphate groups from it. [35]

    . CD45 is mainly produced in blood cells. [4] In humans it has been shown to be able to act on JAK1 and JAK3, [48] whereas in mice, CD45 is capable of acting on all JAKs. [49] One study indicates that CD45 can reduce the amount of time that JAK-STAT signalling is active. [49] The exact details of how CD45 functions is still unknown. [35]

Suppressors of cytokine signalling (SOCS) Edit

There are eight protein members of the SOCS family: cytokine-inducible SH2 domain-containing protein (CISH), SOCS1, SOCS2, SOCS3, SOCS4, SOCS5, SOCS6, and SOCS7, each protein has an SH2 domain and a 40-amino-acid region called the SOCS box. [50] The SOCS box can interact with a number of proteins to form a protein complex, and this complex can then cause the breakdown of JAKs and the receptors themselves, therefore inhibiting JAK-STAT signalling. [4] The protein complex does this by allowing a marker called ubiquitin to be added to proteins, in a process called ubiquitination, which signals for a protein to be broken down. [51] The proteins, such as JAKs and the receptors, are then transported to a compartment in the cell called the proteasome, which carries out protein breakdown. [51]

SOCS can also function by binding to proteins involved in JAK-STAT signalling and blocking their activity. For example, the SH2 domain of SOCS1 binds to a tyrosine in the activation loop of JAKs, which prevents JAKs from phosphorylating each other. [4] The SH2 domains of SOCS2, SOCS3 and CIS bind directly to receptors themselves. [51] Also, SOCS1 and SOCS3 can prevent JAK-STAT signalling by binding to JAKs, using segments called kinase inhibitory regions (KIRs) and stopping JAKs binding to other proteins. [52] The exact details of how other SOCS function is less understood. [4]

Regulator Positive or Negative regulation Function
PTPs SHP-1 and SHP-2: Negative, but could also be positive. CD45, PTP1B, TC-PTP: Negative Removes phosphate groups from receptors, JAKs and STATs
SOCS Negative SOCS1 and SOCS3 block JAKs active sites using KIR domains. SOCS2, SOCS3 and CIS can bind receptors. SOCS1 and SOCS3 can signal JAKs and receptor for degradation.
PIAS Negative Add SUMO group to STATs to inhibit STAT activity. Recruit histone deacetylases to lower gene expression. Prevent STATs binding to DNA.

Since the JAK-STAT pathway plays a major role in many fundamental processes, such as apoptosis and inflammation, dysfunctional proteins in the pathway may lead to a number of diseases. For example, alterations in JAK-STAT signalling can result in cancer and diseases affecting the immune system, such as severe combined immunodeficiency disorder (SCID). [53]

Immune system-related diseases Edit

JAK3 can be used for the signalling of IL-2, IL-4, IL-15 and IL-21 (as well as other cytokines) therefore patients with mutations in the JAK3 gene often experience issues affecting many aspects of the immune system. [54] [55] For example, non-functional JAK3 causes SCID, which results in patients having no NK cells, B cells or T cells, and this would make SCID individuals susceptible to infection. [55] Mutations of the STAT5 protein, which can signal with JAK3, has been shown to result in autoimmune disorders. [56]

It has been suggested that patients with mutations in STAT1 and STAT2 are often more likely to develop infections from bacteria and viruses. [57] Also, STAT4 mutations have been associated with rheumatoid arthritis, and STAT6 mutations are linked to asthma. [58] [59]

Patients with a faulty JAK-STAT signalling pathway may also experience skin disorders. For example, non-functional cytokine receptors, and overexpression of STAT3 have both been associated with psoriasis (an autoimmune disease associated with red, flaky skin). [55] STAT3 plays an important role in psoriasis, as STAT3 can control the production of IL-23 receptors, and IL-23 can help the development of Th17 cells, and Th17 cells can induce psoriasis. [60] Also, since many cytokines function through the STAT3 transcription factor, STAT3 plays a significant role in maintaining skin immunity. [55] In addition, because patients with JAK3 gene mutations have no functional T cells, B cells or NK cells, they would more likely to develop skin infections.

Cancer Edit

Cancer involves abnormal and uncontrollable cell growth in a part of the body. Therefore, since JAK-STAT signalling can allow the transcription of genes involved in cell division, one potential effect of excessive JAK-STAT signalling is cancer formation. High levels of STAT activation have been associated with cancer in particular, high amounts of STAT3 and STAT5 activation is mostly linked to more dangerous tumours. [61] For example, too much STAT3 activity has been associated with increasing the likelihood of melanoma (skin cancer) returning after treatment and abnormally high levels of STAT5 activity have been linked to a greater probability of patient death from prostate cancer. [62] [61] Altered JAK-STAT signalling can also be involved in developing breast cancer. JAK-STAT signalling in mammary glands (located within breasts) can promote cell division and reduce cell apoptosis during pregnancy and puberty, and therefore if excessively activated, cancer can form. [63] High STAT3 activity plays a major role in this process, as it can allow the transcription of genes such as BCL2 and c-Myc, which are involved in cell division. [63]

Mutations in JAK2 can lead to leukaemia and lymphoma. [6] Specifically, mutations in exons 12, 13, 14 and 15 of the JAK2 gene are proposed to be a risk factor in developing lymphoma or leukemia. [6] Additionally, mutated STAT3 and STAT5 can increase JAK-STAT signalling in NK and T cells, which promotes very high proliferation of these cells, and increases the likelihood of developing leukaemia. [63] Also, a JAK-STAT signalling pathway mediated by erythropoietin (EPO), which usually allows the development of red blood cells, may be altered in patients with leukemia. [64]

Treatments Edit

Since excessive JAK-STAT signalling is responsible for some cancers and immune disorders, JAK inhibitors have been proposed as drugs for therapy. For instance, to treat some forms of leukaemia, targeting and inhibiting JAKs could eliminate the effects of EPO signalling and perhaps prevent the development of leukaemia. [64] One example of a JAK inhibitor drug is Ruxolitinib, which is used as a JAK2 inhibitor. [61] STAT inhibitors are also being developed, and many of the inhibitors target STAT3. [63] It has been reported that therapies which target STAT3 can improve the survival of patients with cancer. [63] Another drug, called Tofacitinib, has been used for psoriasis and rheumatoid arthritis treatment, and has been recently approved for Crohn's Disease and Ulcerative Colitis treatment. [53]

Selected Reviews:

  • Gilmore TD (2008)
  • Hayden MS, Ghosh S (2008) Shared principles in NF-kappaB signaling. Cell 132(3), 344–62.
  • Perkins ND (2006) Post-translational modifications regulating the activity and function of the nuclear factor kappa B pathway. Oncogene 25(51), 6717–30.
  • Sun SC (2012) The noncanonical NF-κB pathway. Immunol. Rev. 246(1), 125–40.
  • Chen J, Chen ZJ (2013) Regulation of NF-κB by ubiquitination. Curr. Opin. Immunol. 25(1), 4–12.

We would like to thank Prof. Thomas D. Gilmore, Boston University, Boston, MA, for contributing to this diagram.


The identification of erythropoietin receptors (EpoR) on cancer cells has caused concern, since it implies the possibility that treatment of cancer patients with erythropoietin (Epo) and related agents with demonstrable antiapoptotic activity could enhance cancer growth and progression. However, the function and even the validity of the identification of these receptors have been called into question. We now report the characterization of EpoR and Epo expression by 4 human ovarian cancer cell lines: A2780, CaOV, SKOV and OVCAR-3. Using semiquantitative RT-PCR, restriction digestion of the PCR products and DNA sequence analysis, we determined that each of the lines expresses the EpoR and Epo at the mRNA level. A2780 cells were the highest expressers of both genes. We demonstrated EpoR protein both by western blotting and by immunofluorescence and biologically active Epo protein by quantitative in vitro bioassay. The EpoR on A2780 cells was shown to be functional, since Epo stimulation resulted in phosphorylation of Erk1/2, an important EpoR mitogenic signaling intermediate. None of the cell lines exhibited a growth response in culture to exogenous Epo. However, addition of a neutralizing anti-Epo antibody to A2780 cells resulted in partial growth inhibition that was reversed by the addition of excess Epo, providing evidence for an autocrine/paracrine mechanism of growth enhancement in these cells. © 2007 Wiley-Liss, Inc.

Erythropoietin (Epo) is a glycoprotein hormone that functions as both a growth/survival factor and regulator of differentiation during red blood cell development. In recent years, Epo has been shown to act on nonhematopoietic cells, such as the central nervous system, the heart and other organs and tissues resulting in “tissue protection” 1 from a variety of insults (reviewed in Refs. 2 and 3 ). The erythropoietin receptor (EpoR) has been reportedly identified on numerous normal and neoplastic tissues. However, in many instances, this identification was made solely with a single, commercially available antibody, which has now been proven to exhibit nonspecific binding to one or more nonreceptor proteins, thus calling into question many of these reports. 4 Additionally, clear examples of Epo-mediated EpoR signaling events in these cells are rare. 5

Both Epo and EpoR have been identified in the female reproductive tract, including the ovary, where Epo apparently is produced locally under estrogen regulation. 6 , 7 There is experimental evidence that interruption of Epo/EpoR signaling either in vitro or in vivo can lead to shrinkage of ovarian tumors and their accompanying vasculature. 8 , 9

We now report the characterization of EpoR and Epo expression by 4 human ovarian cancer cell lines. We demonstrate that each line expresses both EpoR and Epo at the mRNA and protein levels and also show signaling from the EpoR in one of the lines. Furthermore, addition of a neutralizing anti-Epo monoclonal antibody reduced growth/viability of the cells. These observations are consistent with the existence of an Epo/EpoR autocrine and/or paracrine mechanism in ovarian cancer.


Borchmann P, Eichenauer DA, Engert A. State of the art in the treatment of Hodgkin lymphoma. Nat Rev Clin Oncol. 20129:450–9.

Younes A, Ansell SM. Novel agents in the treatment of Hodgkin lymphoma: biological basis and clinical results. Semin Hematol. 201653:186–9.

Swerdlow SH, Campo E, Pileri SA, Harris NL, Stein H, Siebert R, et al. The 2016 revision of the World Health Organization classification of lymphoid neoplasms. Blood. 2016127:2375–90.

Hodgkin T. On some morbid appearances of the absorbent glands and spleen. Med Chir Trans. 183217:68–114.

Foss HD, Reusch R, Demel G, Lenz G, Anagnostopoulos I, Hummel M, et al. Frequent expression of the B-cell-specific activator protein in Reed-Sternberg cells of classical Hodgkin’s disease provides further evidence for its B-cell origin. Blood. 199994:3108–13.

Schwering I, Bräuninger A, Klein U, Jungnickel B, Tinguely M, Diehl V, et al. Loss of the B-lineage-specific gene expression program in Hodgkin and Reed-Sternberg cells of Hodgkin lymphoma. Blood. 2003101:1505–12.

Tiacci E, Döring C, Brune V, van Noesel CJ, Klapper W, Mechtersheimer G, et al. Analyzing primary Hodgkin and Reed-Sternberg cells to capture the molecular and cellular pathogenesis of classical Hodgkin lymphoma. Blood. 2012120:4609–20.

Bräuninger A, Wacker HH, Rajewsky K, Küppers R, Hansmann ML. Typing the histogenetic origin of the tumor cells of lymphocyte-rich classical Hodgkin’s lymphoma in relation to tumor cells of classical and lymphocyte-predominance Hodgkin’s lymphoma. Cancer Res. 200363:1644–51.

Kanzler H, Küppers R, Hansmann ML. Rajewsky K. Hodgkin and Reed-Sternberg cells in Hodgkin’s disease represent the outgrowth of a dominant tumor clone derived from (crippled) germinal center B cells. J Exp Med. 1996184:1495–505.

Küppers R, Rajewsky K, Zhao M, Simons G, Laumann R, Fischer R, et al. Hodgkin disease: Hodgkin and Reed-Sternberg cells picked from histological sections show clonal immunoglobulin gene rearrangements and appear to be derived from B cells at various stages of development. Proc Natl Acad Sci USA. 199491:10962–6.

Marafioti T, Hummel M, Foss H-D, Laumen H, Korbjuhn P, Anagnostopoulos I, et al. Hodgkin and Reed-Sternberg cells represent an expansion of a single clone originating from a germinal center B-cell with functional immunoglobulin gene rearrangements but defective immunoglobulin transcription. Blood. 200095:1443–50.

Küppers R, Engert A, Hansmann M-L. Hodgkin lymphoma. J Clin Invest. 2012122:3439–47.

Müschen M, Rajewsky K, Bräuninger A, Baur AS, Oudejans JJ, Roers A, et al. Rare occurrence of classical Hodgkin’s disease as a T cell lymphoma. J Exp Med. 2000191:387–94.

Seitz V, Hummel M, Marafioti T, Anagnostopoulos I, Assaf C, Stein H. Detection of clonal T-cell receptor gamma-chain gene rearrangements in Reed-Sternberg cells of classic Hodgkin disease. Blood. 200095:3020–4.

Willenbrock K, Küppers R, Renne C, Brune V, Eckerle S, Weidmann E, et al. Common features and differences in the transcriptome of large cell anaplastic lymphoma and classical Hodgkin’s lymphoma. Haematologica. 200691:596–604.

Weniger MA, Tiacci E, Schneider S, Arnolds J, Rüschenbaum S, Duppach J, et al. Human CD30+ B cells represent a unique subset related to Hodgkin lymphoma cells. J Clin Invest. 2018128:2996–3007.

Bräuninger A, Schmitz R, Bechtel D, Renne C, Hansmann ML, Küppers R. Molecular biology of Hodgkin’s and Reed/Sternberg cells in Hodgkin’s lymphoma. Int J Cancer. 2006118:1853–61.

Kapatai G, Murray P. Contribution of the Epstein Barr virus to the molecular pathogenesis of Hodgkin lymphoma. J Clin Pathol. 200760:1342–9.

Mancao C, Hammerschmidt W. Epstein-Barr virus latent membrane protein 2A is a B-cell receptor mimic and essential for B-cell survival. Blood. 2007110:3715–21.

Greiner A, Tobollik S, Buettner M, Jungnickel B, Herrmann K, Kremmer E, et al. Differential expression of activation-induced cytidine deaminase (AID) in nodular lymphocyte-predominant and classical Hodgkin lymphoma. J Pathol. 2005205:541–7.

Küppers R. The biology of Hodgkin’s lymphoma. Nat Rev Cancer. 20099:15–27.

Braeuninger A, Küppers R, Strickler JG, Wacker HH, Rajewsky K, Hansmann ML. Hodgkin and Reed-Sternberg cells in lymphocyte predominant Hodgkin disease represent clonal populations of germinal center-derived tumor B cells. Proc Natl Acad Sci USA. 199794:9337–42.

Marafioti T, Hummel M, Anagnostopoulos I, Foss HD, Falini B, Delsol G, et al. Origin of nodular lymphocyte-predominant Hodgkin’s disease from a clonal expansion of highly mutated germinal-center B cells. N Engl J Med. 1997337:453–8.

Brune V, Tiacci E, Pfeil I, Döring C, Eckerle S, van Noesel CJM, et al. Origin and pathogenesis of nodular lymphocyte-predominant Hodgkin lymphoma as revealed by global gene expression analysis. J Exp Med. 2008205:2251–68.

Thurner L, Hartmann S, Fadle N, Regitz E, Kemele M, Kim YJ, et al. Lymphocyte predominant cells detect Moraxella catarrhalis-derived antigens in nodular lymphocyte-predominant Hodgkin lymphoma. Nat Commun. 202011:2465.

Weber-Matthiesen K, Deerberg J, Poetsch M, Grote W, Schlegelberger B. Numerical chromosome aberrations are present within the CD30+ Hodgkin and Reed-Sternberg cells in 100% of analyzed cases of Hodgkin’s disease. Blood. 199586:1464–8.

Martin-Subero JI, Klapper W, Sotnikova A, Callet-Bauchu E, Harder L, Bastard C, et al. Chromosomal breakpoints affecting immunoglobulin loci are recurrent in Hodgkin and Reed-Sternberg cells of classical Hodgkin lymphoma. Cancer Res. 200666:10332–8.

Cuceu C, Hempel WM, Sabatier L, Bosq J, Carde P, M’Kacher R. Chromosomal instability in Hodgkin lymphoma: an in-depth review and perspectives. Cancers (Basel). 201810:91.

Rengstl B, Newrzela S, Heinrich T, Weiser C, Thalheimer FB, Schmid F, et al. Incomplete cytokinesis and re-fusion of small mononucleated Hodgkin cells lead to giant multinucleated Reed-Sternberg cells. Proc Natl Acad Sci USA. 2013110:20729–34.

Ikeda J, Mamat S, Tian T, Wang Y, Rahadiani N, Aozasa K, et al. Tumorigenic potential of mononucleated small cells of Hodgkin lymphoma cell lines. Am J Pathol. 2010177:3081–8.

Reichel J, Chadburn A, Rubinstein PG, Giulino-Roth L, Tam W, Liu Y, et al. Flow-sorting and exome sequencing reveals the oncogenome of primary Hodgkin and Reed-Sternberg cells. Blood. 2015125:1061–72.

Tiacci E, Ladewig E, Schiavoni G, Penson A, Fortini E, Pettirossi V, et al. Pervasive mutations of JAK-STAT pathway genes in classical Hodgkin lymphoma. Blood. 2018131:2454–65.

Wienand K, Chapuy B, Stewart C, Dunford AJ, Wu D, Kim J, et al. Genomic analyses of flow-sorted Hodgkin Reed-Sternberg cells reveal complementary mechanisms of immune evasion. Blood Adv. 20193:4065–80.

Weniger MA, Küppers R. NF-kappaB deregulation in Hodgkin lymphoma. Semin Cancer Biol. 201639:32–9.

Joos S, Menz CK, Wrobel G, Siebert R, Gesk S, Ohl S, et al. Classical Hodgkin lymphoma is characterized by recurrent copy number gains of the short arm of chromosome 2. Blood. 200299:1381–7.

Martin-Subero JI, Gesk S, Harder L, Sonoki T, Tucker PW, Schlegelberger B, et al. Recurrent involvement of the REL and BCL11A loci in classical Hodgkin lymphoma. Blood. 200299:1474–7.

Martin-Subero JI, Wlodarska I, Bastard C, Picquenot JM, Höppner J, Giefing M, et al. Chromosomal rearrangements involving the BCL3 locus are recurrent in classical Hodgkin and peripheral T-cell lymphoma. Blood. 2006108:401–2.

Steidl C, Telenius A, Shah SP, Farinha P, Barclay L, Boyle M, et al. Genome-wide copy number analysis of Hodgkin Reed-Sternberg cells identifies recurrent imbalances with correlations to treatment outcome. Blood. 2010116:418–27.

Jungnickel B, Staratschek-Jox A, Bräuninger A, Spieker T, Wolf J, Diehl V, et al. Clonal deleterious mutations in the IkappaBalpha gene in the malignant cells in Hodgkin’s lymphoma. J Exp Med. 2000191:395–402.

Schmitz R, Hansmann ML, Bohle V, Martin-Subero JI, Hartmann S, Mechtersheimer G, et al. TNFAIP3 (A20) is a tumor suppressor gene in Hodgkin lymphoma and primary mediastinal B cell lymphoma. J Exp Med. 2009206:981–9.

Emmerich F, Theurich S, Hummel M, Haeffker A, Vry MS, Döhner K, et al. Inactivating I kappa B epsilon mutations in Hodgkin/Reed-Sternberg cells. J Pathol. 2003201:413–20.

Kato M, Sanada M, Kato I, Sato Y, Takita J, Takeuchi K, et al. Frequent inactivation of A20 in B-cell lymphomas. Nature. 2009459:712–6.

Otto C, Giefing M, Massow A, Vater I, Gesk S, Schlesner M, et al. Genetic lesions of the TRAF3 and MAP3K14 genes in classical Hodgkin lymphoma. Br J Haematol. 2012157:702–8.

Schmidt A, Schmitz R, Giefing M, Martin-Subero JI, Gesk S, Vater I, et al. Rare occurrence of biallelic CYLD gene mutations in classical Hodgkin lymphoma. Genes Chromosomes Cancer. 201049:803–9.

Lake A, Shield LA, Cordano P, Chui DT, Osborne J, Crae S, et al. Mutations of NFKBIA, encoding IkappaB alpha, are a recurrent finding in classical Hodgkin lymphoma but are not a unifying feature of non-EBV-associated cases. Int J Cancer. 2009125:1334–42.

Gunawardana J, Chan FC, Telenius A, Woolcock B, Kridel R, Tan KL, et al. Recurrent somatic mutations of PTPN1 in primary mediastinal B cell lymphoma and Hodgkin lymphoma. Nat Genet. 201446:329–35.

Weniger MA, Melzner I, Menz CK, Wegener S, Bucur AJ, Dorsch K, et al. Mutations of the tumor suppressor gene SOCS-1 in classical Hodgkin lymphoma are frequent and associated with nuclear phospho-STAT5 accumulation. Oncogene. 200625:2679–84.

Joos S, Küpper M, Ohl S, von Bonin F, Mechtersheimer G, Bentz M, et al. Genomic imbalances including amplification of the tyrosine kinase gene JAK2 in CD30+ Hodgkin cells. Cancer Res. 200060:549–52.

Rui L, Emre NC, Kruhlak MJ, Chung HJ, Steidl C, Slack G, et al. Cooperative epigenetic modulation by cancer amplicon genes. Cancer Cell. 201018:590–605.

Hartmann S, Martin-Subero JI, Gesk S, Husken J, Giefing M, Nagel I, et al. Detection of genomic imbalances in microdissected Hodgkin and Reed-Sternberg cells of classical Hodgkin’s lymphoma by array-based comparative genomic hybridization. Haematologica. 200893:1318–26.

Desch AK, Hartung K, Botzen A, Brobeil A, Rummel M, Kurch L, et al. Genotyping circulating tumor DNA of pediatric Hodgkin lymphoma. Leukemia. 202034:151–66.

Green MR, Monti S, Rodig SJ, Juszczynski P, Currie T, O’Donnell E, et al. Integrative analysis reveals selective 9p24.1 amplification, increased PD-1 ligand expression, and further induction via JAK2 in nodular sclerosing Hodgkin lymphoma and primary mediastinal large B-cell lymphoma. Blood. 2010116:3268–77.

Roemer MG, Advani RH, Ligon AH, Natkunam Y, Redd RA, Homer H, et al. PD-L1 and PD-L2 genetic alterations define classical Hodgkin lymphoma and predict outcome. J Clin Oncol. 201634:2690–7.

Steidl C, Shah SP, Woolcock BW, Rui L, Kawahara M, Farinha P, et al. MHC class II transactivator CIITA is a recurrent gene fusion partner in lymphoid cancers. Nature. 2011471:377–81.

Schneider M, Schneider S, Zühlke-Jenisch R, Klapper W, Sundström C, Hartmann S, et al. Alterations of the CD58 gene in classical Hodgkin lymphoma. Genes Chromosomes Cancer. 201554:638–45.

Camus V, Stamatoullas A, Mareschal S, Viailly PJ, Sarafan-Vasseur N, Bohers E, et al. Detection and prognostic value of recurrent exportin 1 mutations in tumor and cell-free circulating DNA of patients with classical Hodgkin lymphoma. Haematologica. 2016101:1094–101.

Salipante SJ, Adey A, Thomas A, Lee C, Liu YJ, Kumar A, et al. Recurrent somatic loss of TNFRSF14 in classical Hodgkin lymphoma. Genes Chromosomes Cancer. 201655:278–87.

Wlodarska I, Nooyen P, Maes B, Martin-Subero JI, Siebert R, Pauwels P, et al. Frequent occurrence of BCL6 rearrangements in nodular lymphocyte predominance Hodgkin lymphoma but not in classical Hodgkin lymphoma. Blood. 2003101:706–10.

Schumacher MA, Schmitz R, Brune V, Tiacci E, Döring C, Hansmann ML, et al. Mutations in the genes coding for the NF-kappaB regulating factors IkappaBalpha and A20 are uncommon in nodular lymphocyte-predominant Hodgkin’s lymphoma. Haematologica. 201095:153–7.

Hartmann S, Döring C, Vucic E, Chan FC, Ennishi D, Tousseyn T, et al. Array comparative genomic hybridization reveals similarities between nodular lymphocyte predominant Hodgkin lymphoma and T cell/histiocyte rich large B cell lymphoma. Br J Haematol. 2015169:415–22.

Mottok A, Renné C, Willenbrock K, Hansmann ML, Bräuninger A. Somatic hypermutation of SOCS1 in lymphocyte-predominant Hodgkin lymphoma is accompanied by high JAK2 expression and activation of STAT6. Blood. 2007110:3387–90.

Hartmann S, Schuhmacher B, Rausch T, Fuller L, Döring C, Weniger M, et al. Highly recurrent mutations of SGK1, DUSP2 and JUNB in nodular lymphocyte predominant Hodgkin lymphoma. Leukemia. 201630:844–53.

Morin RD, Mendez-Lago M, Mungall AJ, Goya R, Mungall KL, Corbett RD, et al. Frequent mutation of histone-modifying genes in non-Hodgkin lymphoma. Nature. 2011476:298–303.

Lollies A, Hartmann S, Schneider M, Bracht T, Weiss AL, Arnolds J, et al. An oncogenic axis of STAT-mediated BATF3 upregulation causing MYC activity in classical Hodgkin lymphoma and anaplastic large cell lymphoma. Leukemia. 201832:92–101.

Mathas S, Hinz M, Anagnostopoulos I, Krappmann D, Lietz A, Jundt F, et al. Aberrantly expressed c-Jun and JunB are a hallmark of Hodgkin lymphoma cells, stimulate proliferation and synergize with NF-kappa B. EMBO J. 200221:4104–13.

Bargou RC, Emmerich F, Krappmann D, Bommert K, Mapara MY, Arnold W, et al. Constitutive nuclear factor-kappaB-RelA activation is required for proliferation and survival of Hodgkin’s disease tumor cells. J Clin Invest. 1997100:2961–9.

Carbone A, Gloghini A, Gruss HJ, Pinto A. CD40 ligand is constitutively expressed in a subset of T cell lymphomas and on the microenvironmental reactive T cells of follicular lymphomas and Hodgkin’s disease. Am J Pathol. 1995147:912–22.

Molin D, Fischer M, Xiang Z, Larsson U, Harvima I, Venge P, et al. Mast cells express functional CD30 ligand and are the predominant CD30L-positive cells in Hodgkin’s disease. Br J Haematol. 2001114:616–23.

Hirsch B, Hummel M, Bentink S, Fouladi F, Spang R, Zollinger R, et al. CD30-induced signaling is absent in Hodgkin’s cells but present in anaplastic large cell lymphoma cells. Am J Pathol. 2008172:510–20.

Horie R, Watanabe T, Morishita Y, Ito K, Ishida T, Kanegae Y, et al. Ligand-independent signaling by overexpressed CD30 drives NF-kappaB activation in Hodgkin-Reed-Sternberg cells. Oncogene. 200221:2493–503.

Kilger E, Kieser A, Baumann M, Hammerschmidt W. Epstein-Barr virus-mediated B-cell proliferation is dependent upon latent membrane protein 1, which simulates an activated CD40 receptor. EMBO J. 199817:1700–9.

de Oliveira KAP, Kaergel E, Heinig M, Fontaine J-F, Patone G, Muro EM, et al. A roadmap of constitutive NF-kB activity in Hodgkin lymphoma: Dominant roles of p50 and p52 revealed by genome-wide analyses. Genome Med. 20168:28.

Kapp U, Yeh WC, Patterson B, Elia AJ, Kagi D, Ho A, et al. Interleukin 13 is secreted by and stimulates the growth of Hodgkin and Reed-Sternberg cells. J Exp Med. 1999189:1939–46.

Lamprecht B, Kreher S, Anagnostopoulos I, Johrens K, Monteleone G, Jundt F, et al. Aberrant expression of the Th2 cytokine IL-21 in Hodgkin lymphoma cells regulates STAT3 signaling and attracts Treg cells via regulation of MIP-3. Blood. 2008112:3339–47.

Scheeren FA, Diehl SA, Smit LA, Beaumont T, Naspetti M, Bende RJ, et al. IL-21 is expressed in Hodgkin lymphoma and activates STAT5 evidence that activated STAT5 is required for Hodgkin lymphomagenesis. Blood. 2008111:4706–15.

Cattaruzza L, Gloghini A, Olivo K, Di Francia R, Lorenzon D, De Filippi R, et al. Functional coexpression of Interleukin (IL)-7 and its receptor (IL-7R) on Hodgkin and Reed-Sternberg cells: Involvement of IL-7 in tumor cell growth and microenvironmental interactions of Hodgkin’s lymphoma. Int J Cancer. 2009125:1092–101.

Kube D, Holtick U, Vockerodt M, Ahmadi T, Behrmann I, Heinrich PC, et al. STAT3 is constitutively activated in Hodgkin cell lines. Blood. 200198:762–70.

Skinnider BF, Elia AJ, Gascoyne RD, Patterson B, Trümper L, Kapp U, et al. Signal transducer and activator of transcription 6 is frequently activated in Hodgkin and Reed-Sternberg cells of Hodgkin lymphoma. Blood. 200299:618–26.

Dominguez-Sola D, Kung J, Holmes AB, Wells VA, Mo T, Basso K, et al. The FOXO1 transcription factor instructs the germinal center dark zone program. Immunity. 201543:1064–74.

Sander S, Chu VT, Yasuda T, Franklin A, Graf R, Calado DP, et al. PI3 kinase and FOXO1 transcription factor activity differentially control B cells in the germinal center light and dark zones. Immunity. 201543:1075–86.

Dutton A, Reynolds GM, Dawson CW, Young LS, Murray PG. Constitutive activation of phosphatidyl-inositide 3 kinase contributes to the survival of Hodgkin’s lymphoma cells through a mechanism involving Akt kinase and mTOR. J Pathol. 2005205:498–506.

Xie L, Ushmorov A, Leithäuser F, Guan H, Steidl C, Farbinger J, et al. FOXO1 is a tumor suppressor in classical Hodgkin lymphoma. Blood. 2012119:3503–11.

Vrzalikova K, Ibrahim M, Vockerodt M, Perry T, Margielewska S, Lupino L, et al. S1PR1 drives a feedforward signalling loop to regulate BATF3 and the transcriptional programme of Hodgkin lymphoma cells. Leukemia. 201832:214–23.

Muppidi JR, Schmitz R, Green JA, Xiao W, Larsen AB, Braun SE, et al. Loss of signalling via Galpha13 in germinal centre B-cell-derived lymphoma. Nature. 2014516:254–8.

Renné C, Willenbrock K, Küppers R, Hansmann M-L, Bräuninger A. Autocrine and paracrine activated receptor tyrosine kinases in classical Hodgkin lymphoma. Blood. 2005105:4051–9.

Teofili L, Di Febo AL, Pierconti F, Maggiano N, Bendandi M, Rutella S, et al. Expression of the c-met proto-oncogene and its ligand, hepatocyte growth factor, in Hodgkin disease. Blood. 200197:1063–9.

Lamprecht B, Walter K, Kreher S, Kumar R, Hummel M, Lenze D, et al. Derepression of an endogenous long terminal repeat activates the CSF1R proto-oncogene in human lymphoma. Nat Med. 201016:571–9.

Cader FZ, Vockerodt M, Bose S, Nagy E, Brundler MA, Kearns P, et al. The EBV oncogene LMP1 protects lymphoma cells from cell death through the collagen-mediated activation of DDR1. Blood. 2013122:4237–45.

Moreau A, Praloran V, Berrada L, Coupey L, Gaillard F. Immunohistochemical detection of cells positive for colony-stimulating factor 1 in lymph nodes from reactive lymphadenitis, and Hodgkin’s disease. Leukemia. 19926:126–30.

Edginton-White B, Cauchy P, Assi SA, Hartmann S, Riggs AG, Mathas S, et al. Global long terminal repeat activation participates in establishing the unique gene expression programme of classical Hodgkin lymphoma. Leukemia. 201933:1463–74.

Renne C, Minner S, Küppers R, Hansmann ML, Bräuninger A. Autocrine NGFbeta/TRKA signalling is an important survival factor for Hodgkin lymphoma derived cell lines. Leuk Res. 200832:163–7.

Zheng B, Fiumara P, Li YV, Georgakis G, Snell V, Younes M, et al. MEK/ERK pathway is aberrantly active in Hodgkin disease: a signaling pathway shared by CD30, CD40, and RANK that regulates cell proliferation and survival. Blood. 2003102:1019–27.

Juszczynski P, Ouyang J, Monti S, Rodig SJ, Takeyama K, Abramson J, et al. The AP1-dependent secretion of galectin-1 by Reed Sternberg cells fosters immune privilege in classical Hodgkin lymphoma. Proc Natl Acad Sci USA. 2007104:13134–9.

Watanabe M, Ogawa Y, Ito K, Higashihara M, Kadin ME, Abraham LJ, et al. AP-1 mediated relief of repressive activity of the CD30 promoter microsatellite in Hodgkin and Reed-Sternberg cells. Am J Pathol. 2003163:633–41.

Jundt F, Anagnostopoulos I, Förster R, Mathas S, Stein H, Dörken B. Activated Notch 1 signaling promotes tumor cell proliferation and survival in Hodgkin and anaplastic large cell lymphoma. Blood. 200299:3398–403.

Aldinucci D, Lorenzon D, Cattaruzza L, Pinto A, Gloghini A, Carbone A, et al. Expression of CCR5 receptors on Reed-Sternberg cells and Hodgkin lymphoma cell lines: involvement of CCL5/Rantes in tumor cell growth and microenvironmental interactions. Int J Cancer. 2008122:769–76.

Skinnider BF, Mak TW. The role of cytokines in classical Hodgkin lymphoma. Blood. 200299:4283–97.

van den Berg A, Visser L, Poppema S. High expression of the CC chemokine TARC in Reed-Sternberg cells. A possible explanation for the characteristic T-cell infiltration Hodgkin’s lymphoma. Am J Pathol. 1999154:1685–91.

Hansen HP, Engels HM, Dams M, Paes Leme AF, Pauletti BA, Simhadri VL, et al. Protrusion-guided extracellular vesicles mediate CD30 trans-signalling in the microenvironment of Hodgkin’s lymphoma. J Pathol. 2014232:405–14.

Aoki T, Chong LC, Takata K, Milne K, Hav M, Colombo A, et al. Single-cell transcriptome analysis reveals disease-defining T-cell subsets in the tumor microenvironment of classic Hodgkin lymphoma. Cancer Discov. 202010:406–21.

Wein F, Weniger MA, Hoing B, Arnolds J, Hüttmann A, Hansmann ML, et al. Complex immune evasion strategies in classical Hodgkin lymphoma. Cancer Immunol Res. 20175:1122–32.

Biggar RJ, Jaffe ES, Goedert JJ, Chaturvedi A, Pfeiffer R, Engels EA. Hodgkin lymphoma and immunodeficiency in persons with HIV/AIDS. Blood. 2006108:3786–91.

Wein F, Küppers R. The role of T cells in the microenvironment of Hodgkin lymphoma. J Leukoc Biol. 201699:45–50.

Ma Y, Visser L, Blokzijl T, Harms G, Atayar C, Poppema S, et al. The CD4+CD26- T-cell population in classical Hodgkin’s lymphoma displays a distinctive regulatory T-cell profile. Lab Invest. 200888:482–90.

Roemer MGM, Redd RA, Cader FZ, Pak CJ, Abdelrahman S, Ouyang J, et al. Major histocompatibility complex class II and programmed death ligand 1 expression predict outcome after programmed death 1 blockade in classic Hodgkin lymphoma. J Clin Oncol. 201836:942–50.

Bosshart H, Jarrett RF. Deficient major histocompatibility complex class II antigen presentation in a subset of Hodgkin’s disease tumor cells. Blood. 199892:2252–9.

Greaves P, Clear A, Owen A, Iqbal S, Lee A, Matthews J, et al. Defining characteristics of classical Hodgkin lymphoma microenvironment T-helper cells. Blood. 2013122:2856–63.

Poppema S. Immunobiology and pathophysiology of Hodgkin lymphomas. Hematology Am Soc Hematol Educ Program. 2005:231–8.

Cader FZ, Schackmann RCJ, Hu X, Wienand K, Redd R, Chapuy B, et al. Mass cytometry of Hodgkin lymphoma reveals a CD4(+) regulatory T-cell-rich and exhausted T-effector microenvironment. Blood. 2018132:825–36.

Lee SP, Constandinou CM, Thomas WA, Croom-Carter D, Blake NW, Murray PG, et al. Antigen presenting phenotype of Hodgkin Reed-Sternberg cells: analysis of the HLA class I processing pathway and the effects of interleukin-10 on epstein-barr virus-specific cytotoxic T-cell recognition. Blood. 199892:1020–30.

Oudejans JJ, Jiwa NM, Kummer JA, Horstman A, Vos W, Baak JP, et al. Analysis of major histocompatibility complex class I expression on Reed-Sternberg cells in relation to the cytotoxic T-cell response in Epstein-Barr virus-positive and -negative Hodgkin’s disease. Blood. 199687:3844–51.

Gandhi MK, Moll G, Smith C, Dua U, Lambley E, Ramuz O, et al. Galectin-1 mediated suppression of Epstein-Barr virus specific T-cell immunity in classic Hodgkin lymphoma. Blood. 2007110:1326–9.

Yamamoto R, Nishikori M, Kitawaki T, Sakai T, Hishizawa M, Tashima M, et al. PD-1-PD-1 ligand interaction contributes to immunosuppressive microenvironment of Hodgkin lymphoma. Blood. 2008111:3220–4.

Vari F, Arpon D, Keane C, Hertzberg MS, Talaulikar D, Jain S, et al. Immune evasion via PD-1/PD-L1 on NK cells and monocyte/macrophages is more prominent in Hodgkin lymphoma than DLBCL. Blood. 2018131:1809–19.

Kawashima M, Carreras J, Higuchi H, Kotaki R, Hoshina T, Okuyama K, et al. PD-L1/L2 protein levels rapidly increase on monocytes via trogocytosis from tumor cells in classical Hodgkin lymphoma. Leukemia. 202034:2405–17.

Ho WT, Pang WL, Chong SM, Castella A, Al-Salam S, Tan TE, et al. Expression of CD137 on Hodgkin and Reed-Sternberg cells inhibits T-cell activation by eliminating CD137 ligand expression. Cancer Res. 201373:652–61.

Ishida T, Ishii T, Inagaki A, Yano H, Komatsu H, Iida S, et al. Specific recruitment of CC chemokine receptor 4-positive regulatory T cells in Hodgkin lymphoma fosters immune privilege. Cancer Res. 200666:5716–22.

Marshall NA, Christie LE, Munro LR, Culligan DJ, Johnston PW, Barker RN, et al. Immunosuppressive regulatory T cells are abundant in the reactive lymphocytes of Hodgkin lymphoma. Blood. 2004103:1755–62.

Gandhi MK, Lambley E, Duraiswamy J, Dua U, Smith C, Elliott S, et al. Expression of LAG-3 by tumor-infiltrating lymphocytes is coincident with the suppression of latent membrane antigen-specific CD8+ T-cell function in Hodgkin lymphoma patients. Blood. 2006108:2280–9.

Patel SS, Weirather JL, Lipschitz M, Lako A, Chen PH, Griffin GK, et al. The microenvironmental niche in classic Hodgkin lymphoma is enriched for CTLA-4-positive T cells that are PD-1-negative. Blood. 2019134:2059–69.

Choe JY, Yun JY, Jeon YK, Kim SH, Park G, Huh JR, et al. Indoleamine 2,3-dioxygenase (IDO) is frequently expressed in stromal cells of Hodgkin lymphoma and is associated with adverse clinical features: a retrospective cohort study. BMC Cancer. 201414:335.

Reinke S, Brockelmann PJ, Iaccarino I, Garcia-Marquez MA, Borchmann S, Jochims F, et al. Tumor and microenvironment response but no cytotoxic T-cell activation in classic Hodgkin lymphoma treated with anti-PD1. Blood. 2020136:2851–63.

Nagasaki J, Togashi Y, Sugawara T, Itami M, Yamauchi N, Yuda J, et al. The critical role of CD4+ T cells in PD-1 blockade against MHC-II-expressing tumors such as classic Hodgkin lymphoma. Blood Adv. 20204:4069–82.

Cader FZ, Hu X, Goh WL, Wienand K, Ouyang J, Mandato E, et al. A peripheral immune signature of responsiveness to PD-1 blockade in patients with classical Hodgkin lymphoma. Nat Med. 202026:1468–79.

Jalali S, Price-Troska T, Bothun C, Villasboas J, Kim HJ, Yang ZZ, et al. Reverse signaling via PD-L1 supports malignant cell growth and survival in classical Hodgkin lymphoma. Blood. Cancer J. 20199:22.

Du J, Neuenschwander M, Yu Y, Dabritz JH, Neuendorff NR, Schleich K, et al. Pharmacological restoration and therapeutic targeting of the B-cell phenotype in classical Hodgkin lymphoma. Blood. 2017129:71–81.

Guan H, Xie L, Wirth T, Ushmorov A. Repression of TCF3/E2A contributes to Hodgkin lymphomagenesis. Oncotarget. 20167:36854–64.

Yuki H, Ueno S, Tatetsu H, Niiro H, Iino T, Endo S, et al. PU.1 is a potent tumor suppressor in classical Hodgkin lymphoma cells. Blood. 2013121:962–70.

Küppers R. Mechanisms of B-cell lymphoma pathogenesis. Nat Rev Cancer. 20055:251–62.

Spina V, Bruscaggin A, Cuccaro A, Martini M, Di Trani M, Forestieri G, et al. Circulating tumor DNA reveals genetics, clonal evolution, and residual disease in classical Hodgkin lymphoma. Blood. 2018131:2413–25.