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Hypoxia. Hypoxia in the pathogenesis of systemic sclerosis

Autoimmunity, microangiopathy and tissue fibrosis are hallmarks of systemic sclerosis (SSc). Vascular alterations and reduced capillary density decrease blood flow and impair tissue oxygenation in SSc. Oxygen supply is further reduced by accumulation of extracellular matrix (ECM), which increases diffusion distances from blood vessels to cells. Therefore, severe hypoxia is a characteristic feature of SSc and might contribute directly to the progression of the disease. Hypoxia stimulates the production of ECM proteins by SSc fibroblasts in a transforming growth factor-β-dependent manner. The induction of ECM proteins by hypoxia is mediated via hypoxia-inducible factor-1α-dependent and -independent pathways. Hypoxia may also aggravate vascular disease in SSc by perturbing vascular endothelial growth factor (VEGF) receptor signalling. Hypoxia is a potent inducer of VEGF and may cause chronic VEGF over-expression in SSc. Uncontrolled over-expression of VEGF has been shown to have deleterious effects on angiogenesis because it leads to the formation of chaotic vessels with decreased blood flow. Altogether, hypoxia might play a central role in pathogenesis of SSc by augmenting vascular disease and tissue fibrosis.

Introduction
Oxygen homeostasis is a sine qua non for metazoan organisms. Reduction in physiological oxygen concentrations leads to metabolic demise because oxygen is the terminal electron acceptor during ATP formation in mitochondria and is a central substrate in many enzymatic reactions. Whereas lack of oxygen causes metabolic cell death, increased oxygen concentrations carry a risk for oxidative damage to proteins, lipids and nucleic acids, possibly initializing apoptosis or carcinogenesis. Thus, even slight changes in systemic and cellular oxygen concentrations induce a tightly regulated machinery of short-acting and long-acting response pathways to keep the supply of oxygen within the physiological range. Molecular responses to hypoxia and endogenous hypoxia markers have been elucidated in detail during the past two decades. In this context, the molecular characterization of the transcription factor hypoxia-inducible factor (HIF)-1 and unravelling of its regulation were breakthroughs for our understanding of cellular adaptation to reduced oxygenation. HIF-1 protein accumulates under hypoxic conditions in many different cell types. It activates the transcription of genes that are of fundamental importance for oxygen homeostasis, including genes involved in energy metabolism, angiogenesis, vasomotor control, apoptosis, proliferation and matrix production [1].
Systemic sclerosis (SSc) is characterized by a triad of microangiopathy, activation of humoral and cellular immune responses and tissue fibrosis, affecting the skin as well as a variety of internal organs, including lung, heart and gastrointestinal tract [2]. Using nailfold capillaroscopy, alterations in the capillary network can be observed early in SSc. Vascular alterations include sac-like, giant and bushy capillaries, microhaemorrhages and a variable loss of capillaries that result in avascular areas [3]. The microangiopathy with progressive loss of capillaries leads to decreased blood flow followed by a lack of nutrients and tissue hypoxia. In advanced disease, fibrosis of the skin and of multiple internal organs, which results from excessive extracellular matrix production of activated fibroblasts, is the most obvious histopathological hallmark of SSc. Because the accumulation of extracellular matrix increases diffusion distances from blood vessels to cells, tissue malnutrition and hypoxia may be aggravated by fibrosis. In summary, severe tissue hypoxia is present in SSc and may even be involved in disease progression.

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The present review presents current knowledge of molecular signalling pathways in response to hypoxia and discusses the role that hypoxia plays in the pathogenesis of SSc.

Molecular structure of hypoxia-inducible factor-1
In 1995, Wang and coworkers cloned the transcription factor HIF-1, based on its ability to bind to the 3' enhancer region of the erythropoietin gene [4]. Structural analysis revealed two subunits: HIF-1α (120 kDa) and HIF-1β (91 to 94 kDa). Both HIF-1 subunits contain a basic helix-loop-helix domain, enabling them to recognize and bind to specific DNA sequences, called HIF-1 DNA binding sites (HBSs), within the regulatory regions of hypoxia-inducible genes. Both proteins are also charactarized by two Per/ARNT/Sim (PAS) regions located at the amino-termini. Using HIF-1α deletion mutants, Jiang and coworkers [5] demonstrated that the helix-loop-helix domain and the PAS-A region of HIF-1α are sufficient for heterodimerization with HIF-1β. The most intriguing structural element of HIF-1α is the oxygen-dependent degradation domain (ODDD), which links HIF-1α to the cellular oxygen sensor. Under normoxic conditions the hydroxylation of two proline residues within the ODDD results in ubiquitinylation and degradation of HIF-1α. In contrast, hydroxylation and degradation of HIF-1α are decreased in hypoxic milieus because oxygen is the critical substrate in hydroxylation reactions. Thus, lack of oxygen leads to HIF-1α accumulation [6].

Stabilization of hypoxia-inducible factor-1α α protein
In contrast to the expression of HIF-1β, that of HIF-1α is tightly controlled by cellular oxygen levels. Cellular HIF-1α is not detectable under normoxic conditions because it is rapidly degraded after translation. After exposure to low oxygen concentrations, levels of HIF-1α increase exponentially. Maximal response is usually reached at oxygen concentrations of about 0.5% .
Hydroxylation of two proline residues within the ODDD (positions 402 and 564) triggers the oxygen-dependent regulation of HIF-1α. This hydroxylation is catalyzed by a family of 2-oxoglutarate dependent dioxygenases called prolyl hydroxylase domains (PHDs) [7]. During the hydroxylation process, PHDs split molecular oxygen and transfer one oxygen atom to one of the proline residues. The second oxygen atom reacts with 2-oxoglutarate, generating succinate and carbon dioxide. The co-substrate ascorbic acid keeps the ferrous ion of the catalytic site in its bivalent state. The ability of PHDs to modify HIF-1α depends on the concentration of its substrate oxygen. Under normoxic conditions, PHDs hydroxylate HIF-1α efficiently, leading to the rapid degradation of the HIF-1α subunit. In contrast, the rate of hydroxylation is reduced at low oxygen levels. Thus, PHDs function as intracellular oxygen sensors and provide the molecular basis for the regulation of HIF-1α protein concentrations by cellular partial pressure of oxygen [8].
Recently, an endoplasmatic prolyl-4-hydroxylase (P4H) with a transmembrane domain, which is more closely related to the collagen prolyl hydroxylases, has also been shown to hydroxylate HIF-1α in vitro [15].
An additional mechanism for the regulation of HIF-1α stability was demonstrated by Jeong and coworkers [16]. Arrest defective (ARD)1, an acetyltransferase, binds directly to the ODDD of HIF-1α in the cytoplasm and acetylates a single lysine residue at position 532. Acetylation of this specific lysine residue favours the interaction of HIF-1α and the E3 ubiquitin ligase complex, and stimulates the degradation of HIF-1α. As shown by vascular endothelial growth factor (VEGF) promoter-driven luciferase reporter gene assays, ARD1 not only destabilizes HIF-1α protein, but it also downregulates its transactivation activity in ARD1-transfected HT1080 human fibrosarcoma cells under hypoxic conditions. Mutation of lysine residue 532 to arginine or application of antisense ARD1 results in stabilization of HIF-1α even under normoxic conditions [16,17]. In contrast, levels of HIF-1α decreased when deacetylation was inhibited. Finally, mRNA and protein levels of ARD-1 are diminished under hypoxia, resulting in less acetylated HIF-1α [16].
Blocking hydroxylation of proline residues 402 and 564 as well as blocking acetylation of lysine 532 have been demonstrated to prevent degradation of HIF-1α under normoxic conditions, thus abolishing the oxygen-dependent regulation of HIF-1α signalling [6,9,16]. These findings suggest that both pathways -hydroxylation and acetylation of HIF-1α -are essential for the physiological regulation of cellular responses to hypoxia.

Upregulation of prolyl hydroxylase domain activity in chronic hypoxia
Interestingly, PHD2 and PHD3 are induced by hypoxia in a HIF-1α-dependent manner, thereby creating a negative feedback loop of HIF-1α signalling [14,18]. In this context, a functional hypoxia-regulated element has been identified in the PHD3 gene, enabling direct regulation of PHD3 by HIF-1. Recently, Ginouvès and coworkers [19] reported increased PHD activity in response to chronic hypoxia. PHD2 and PHD3 protein levels reached a maximum after 24 hours of hypoxia, whereas PHD activity rose steadily for 7 days, indicating that further mechanisms besides induction of PHDs led to increased PHD activity. Consistent with these findings, PHD activity increased with prolonged hypoxia in vivo. Only low PHD activity but high HIF-1α levels were observed in mice exposed to 6 hours of hypoxia at 8% oxygen, whereas PHD activity increased markedly after 24 hours of hypoxia, resulting in a subsequent reduction in HIF-1α. After 24 hours of 8% oxygen, escalation of hypoxia to 6% oxygen concentration for another 2 hours caused a reaccumulation of HIF-1α [19]. Together these findings suggest that HIF-1α is induced in response to hypoxia, accumulates in acute hypoxia and is removed as the activity of PHDs increases in chronic hypoxia.
Ginouvès and coworkers [19] also suggested a mechanism that may lead to augmented PHD activity that is distinct from PHD gene induction. During hypoxia, HIF-1 induces pyruvate dehydrogenase kinase-1, which has been reported to decrease mitochondrial oxygen consumption by inhibiting mitochondrial respiration [20,21]. Inhibition of mitochondrial respiration may increase intracellular oxygen levels and accelerate oxygen-dependent HIF-1α hydroxylation by PHDs [19]. Therefore, augmented PHD activity in chronic hypoxia might create an effective negative feedback loop for HIF-1α signalling. Although this hypothesis must be confirmed with further experiments, separating acute from chronic hypoxia will certainly gain importance for future studies, especially when evaluating HIF-1α or PHDs as possible therapeutic targets for diseases in which hypoxia has been implicated, such as SSc.

Degradation of hypoxia-inducible factor-α α
The rapid degradation of HIF-1α under normoxic conditions is mediated by the von Hippel-Lindau tumour suppressor protein (pVHL) [22]. The β-subunit of pVHL interacts directly with the ODDD of HIF-1α when proline residue(s) 402 and/or 564 are hydroxylated, but not without this modification. pVHL itself is part of the E3 ubiquitin ligase complex. Interaction of prolinehydroxylated HIF-1α with pVHL/E3 ubiquitin ligase complex activates the ubiquitination machinery, thereby promoting degradation of HIF-1α [1,9,23,24]. A similar mechanism of recognition is proposed for the acetylation of the lysine residue 532 [16]. Under hypoxic conditions, the ODDD is neither hydroxylated nor acetylated, pVHL cannot bind and HIF-1α is not ubiquitinated. Thus, degradation of HIF-1α in the proteasome is inhibited and HIF-1α protein accumulates.

Binding of HIF-1 to HIF binding sites, formation of the transcriptional complex and regulation of HIF-1 transactivation
After translocation into the nucleus HIF-1α dimerizes with ARNT/HIF-1β. The HIF-1 heterodimer then binds via its basic helix-loop-helix domain to the HBS within the hypoxiaresponsive element of most hypoxia-regulated genes [25][26][27]. The HBS is essential but not sufficient for HIF-1 gene activation. Besides the HBS, a complete hypoxia-responsive element contains additional binding sites for transcription factors that are not sensitive to hypoxia. These co-stimulatory factors, including cAMP response element binding protein (CREB)-1 of the lactate dehydrogenase A gene [28] or activator protein-1 (AP-1) in the VEGF gene [29], are also required for efficient transcription of oxygen-sensitive genes. Multimerization of HBS can substitute for additional transcription factors in several HIF-regulated genes [30][31][32][33].
For efficient induction of HIF-1-regulated genes, HIF-1 must be activated. Simple blockade of HIF-1α degradation (for example with chemical proteasome inhibitors such as Ncarbobenzoxyl-L-leucinyl-L-leucinyl-L-norvalinal) results in accumulation of HIF-1α but is often not sufficient for transactivation [34]. Two modifications of HIF-1α involved in the regulation of HIF-1α transactivation have been identified: hydroxylation of the carboxyl-terminal transactivation domain and protein phosphorylation by tyrosine kinase receptors.

Other members of the hypoxia-inducible factor family
Two proteins closely related to HIF-1α have been identified and designated HIF-2α and HIF-3α [39,40]. HIF-2α and HIF-3α are both able to dimerize with HIF-1β and bind to HBSs [41,42]. HIF-2α is similar to HIF-1α with regard to its genomic organization, protein structure, dimerization with HIF-1β, DNA binding and transactivation [22,35,43,44]. Moreover, both proteins accumulate under hypoxic conditions [45][46][47]. However, experiments with knockout mice revealed that HIF-1α and HIF-2α could not compensate for loss of the each other [31,48,49]. This finding suggests that the different α subunits of HIF might not be redundant and possess different biological functions.

Hypoxia in systemic sclerosis
Hypoxia and its central mediator HIF-1 control a large variety of different genes. Upregulation of HIF-1 in response to hypoxia regulates erythropoiesis, angiogenesis and glucose metabolism, as well as cell proliferation and apoptosis [1,7]. Using DNA microarray studies on primary pulmonary arterial endothelial cells, Manalo and coworkers [50] observed that a minimum of 2.6% of all human genes were regulated by hypoxia in a HIF-1-dependent manner. In theory, microangiopathy and tissue fibrosis should result in reduced tissue oxygenation and may provoke HIF-1-dependent response to hypoxia. The reduced capillary density and vascular malformations should lead to decreased blood flow with lack of nutrients and oxygen in involved organs in SSc patients [51]. Besides the microangiopathy, tissue fibrosis might further aggravate tissue malnutrition and hypoxia. The progressive accumulation of extracellular matrix proteins such as collagens, fibronectin and glycosaminoglycans [52] increases distances between cells and their supplying vessels and may impair diffusion. Hence, lack of functional capillaries as well as impaired diffusion implicate significant tissue malnutrition and chronic hypoxia in SSc patients ( Figure 1).
Indeed, two studies demonstrated severe hypoxia in lesional, fibrotic skin of SSc patients [53,54]. In both studies, low oxygen levels were only found in lesional skin of SSc patients, whereas the oxygen levels in nonfibrotic skin were not decreased compared with skin of healthy volunteers.
Using a noninvasive transcutaneous technique to measure oxygen levels, Silverstein and coworkers [53] showed that oxygen levels of fibrotic skin were inversely related to skin thickness. The lowest oxygen levels were measured in SSc patients with severely thickened skin. Indirect correlation of oxygen levels with dermal thickness supports the concepts of impaired diffusion due to accumulation of extracellular matrix in lesional skin of SSc patients. Patients suffering from primary Raynaud's disease did not exhibit hypoxic skin, and oxygen levels were similar to those in healthy individuals.
We quantified oxygen levels in the skin of SSc patients by applying an oxygen partial pressure (PO 2 ) histography method, involving introduction of a small polarographic needle electrode directly into the dermis [54]. To exclude systemic influences on local oxygen levels, we determined arterial oxygen saturation, haemoglobin content, blood pressure and heart rate, and patients rested for at least 10 minutes before the experiment. For each patient about 200 single measurements of PO 2 were taken at a predefined area on the dorsal forearm, and an individual PO 2 mean value was determined. Average PO 2 in the skin of healthy individuals was 33.6 ± 4.1 mmHg (4.4 ± 0.5% oxygen per volume), whereas involved skin of SSc patients exhibited significantly decreased oxygen levels, with a mean PO 2 value of 23.7 ± 2.1 mmHg (3.1 ± 0.3%). In contrast, the average PO 2 in nonfibrotic skin of SSc patients did not differ from that in healthy individuals (mean PO 2 37.9 ± 8.6 mmHg, corresponding to 5.0 ± 1.1%).
In summary, both studies demonstrated that hypoxia is a characteristic feature of involved, fibrotic skin of SSc patients. Although cutaneous blood flow, a potential confounding factor, was not determined in any of these studies, the inverse correlation of skin thickness with cutaneous PO 2 suggests that impaired oxygen diffusion due to extracellular matrix accumulation might contribute to tissue hypoxia in SSc.
Despite severely reduced oxygen levels and despite the overexpression of these growth factors, protein levels of HIF-1α in the skin of SSc patients were even below the levels seen in healthy control skin [54]. Skin specimens from SSc patients did not exhibit increased expression of HIF-1α protein by immunohistochemistry. HIF-1α staining was moderate to high Vicious circle of hypoxia and fibrosis in the pathogenesis of SSc. To the upper left is shown nailfold capillaroscopy from a patient with systemic sclerosis (SSc) with capillary rarification and vascular alterations, including sac-like, giant and bushy capillaries. Vasculopathy leads to reduced blood flow and causes tissue hypoxia in SSc. To the upper right is shown a haematoxylin and eosin stained skin section from an experimental mouse fibrosis model with increased skin thickness due to extracellular matrix (ECM) deposition. ECM deposition increases diffusion distances from blood vessels to cells and reduces tissue oxygenation. In the 'vicious circle', shown at the bottom of the figure, tissue hypoxia leads to activation of dermal fibroblasts and upregulation of ECM production. Further ECM deposition aggravates tissue malnutrition and hypoxia. Hypoxia once again stimulates ECM production in dermal fibroblasts.
in the epidermis of healthy individuals, whereas the expression of HIF-1α in SSc patients was restricted to single keratinocytes. HIF-1α protein was not detectable in the dermis of healthy individuals and SSc patients. Moreover, the HIF-1α expression pattern in involved skin in SSc patients did not correlate with upregulated VEGF, one of the main transcriptional targets of HIF-1α [54].
PHD-dependent HIF-1α negative feedback loops in chronic hypoxic conditions might be a plausible explanation for decreased HIF-1α levels in fibrotic skin of SSc patients. Considering the clinical course of SSc, lesional skin in SSc patients can be categorized as a chronically hypoxic tissue. In this context, low HIF-1α levels may be caused by negative HIF-1α feedback loops, even despite severe hypoxia. Increased PHD activity in response to chronic hypoxia [19] might lead to rapid HIF-1α degradation and decreased HIF-1α levels in fibrotic SSc skin. This theory is also supported by studies on the effects of prolonged hypoxia in murine organs. In mice exposed to 6% oxygen, HIF-1α protein reached maximum levels in the brain after 4 to 5 hours but declined afterward, attaining basal normoxic concentrations after 9 to 12 hours. Similar results were obtained for kidney and liver [59].
However, the low levels of HIF-1α in the skin of SSc patients per se do not argue against the persistent activation of oxygen-sensitive pathways in SSc. Marked and persistent upregulation of the oxygen-dependent gene VEGF is observed in lesional SSc skin even in late stages of SSc. Thus, the response to hypoxia appears to persist in chronic states, but might be driven by HIF-1α-independent pathways, for instance HIF-2α and HIF-3α. However, the role played by other members of the HIF family in the pathogenesis of SSc has not yet been investigated in detail.

Insufficient response to hypoxia: dysregulation of angiogenesis in systemic sclerosis
Angiogenesis and vasculogenesis are fundamental mechanisms in improving oxygenation of hypoxic tissue. HIF-1 promotes vascularization by inducing the expression of multiple angiogenic mediators such as VEGF, placental growth factor, angiopoietin 1 and 2, and PDGF-BB [60]. VEGF drives angiogenesis by activating endothelial cells in hypoxic tissue and vasculogenesis by mobilizing and recruiting endothelial progenitor cells [61][62][63]. In addition, VEGF exhibits synergistic angiogenic effects together with PDGF and fibroblast growth factor-2 [64].
Sufficient tissue vascularization depends on strict regulation of VEGF expression. Chronic and uncontrolled overexpression of VEGF induces the formation of chaotic vessels, characterized by glomeruloid and haemangioma-like morphology [65,66]. Dor and coworkers [67] demonstrated in pTET-VEGF 165 /MHCα-tTa transgenic mice, in which VEGF expression can be conditionally switched off in an organ-dependent manner by feeding tetracycline, that time-dependent regulation of VEGF expression was essential for adequate vascularization. Although short-term over-expression of VEGF induced the formation of new mature and functional vessels in adult organs, prolonged exposure to VEGF without subsequently switching off its gene expression by tetracycline resulted in the formation of irregularly shaped, sac-like vessels leading to reduced blood flow. Irregularly shaped, sac-like vessels are reminiscent of the disturbed vessel morphology in SSc [3]. Hence, the microvascular defects in SSc might partly be caused by uncontrolled overexpression of VEGF.
VEGF levels are markedly upregulated in the skin of SSc patients compared with healthy volunteers [54]. As analyzed by in situ hybridization, the mean percentage of epidermal keratinocytes expressing VEGF was significantly increased in SSc patients compared with normal individuals. These findings were consistent with dermal expression levels of VEGF. In contrast, normal individuals did not exhibit VEGF expression in the dermis. VEGF was expressed in most SSc patients in a variety of different dermal cell types, including fibroblasts, endothelial cells and leucocytes [54]. VEGF was induced in dermal SSc fibroblasts in response to hypoxia, but expression levels did not differ significantly between fibroblasts from SSc patients and those from healthy volunteers [54]. However, as the oxygen levels are significantly lower in lesional skin of SSc patients than in control individuals, the induction of VEGF by hypoxia is only operative in SSc patients, but not in normal volunteers. Both receptors for VEGF, namely VEGF receptors 1 and 2, were also overexpressed in the skin of SSc patients. Therefore, enhanced activation of the VEGF/VEGF receptor axis may lead to typical changes in SSc vascularization, causing tissue malnutrition and hypoxia [54]. Because the expression of VEGF is stimulated by hypoxia, one might speculate that the hypoxia could augment vascular disease in SSc by contributing to persistent over-expression of VEGF. However, it remains to be demonstrated that chronic hypoxia alone is indeed sufficient to cause persistent upregulation of VEGF in vivo. Alternatively, the persistent over-expression of VEGF in SSc might also be driven by cytokines. Interleukin-1β, PDGF and TGF-β are all upregulated in SSc and can stimulate the expression of VEGF [54,68,69].

Induction of fibrosis by hypoxia
Microangiopathy with impaired angiogenesis and excessive accumulation of extracellular matrix may cause severe hypoxia in SSc [53,54]. However, what is the exact role played by hypoxia in the pathogenesis of SSc? Is it just the consequence of microangiopathy and fibrosis or does it contribute to the progression of SSc?
DNA microarray studies revealed the first causal links between hypoxia and fibrosis [50]. Manalo and coworkers [50] detected a striking number of genes encoding collagens or collagen-modifying enzymes that were induced in pulmonary endothelial cells after 24 hours at 1% oxygen. These genes included collagen (COL)1A2, COL4A1, COL4A2, COL5A1, COL9A1 and COL18A1, as well as procollagen prolyl hydroxylases (P4HA1 and P4HA2), lysyl oxidase (LOX) and lysyl hydroxylases (procollagen lysyl hydroxylase and procollagen lysyl hydroxylase 2). Similar links between hypoxia and fibrosis have also been found in other models and organs, for example kidney [70,71], liver [72] and lung [73]. Together, these findings indicate that hypoxia could promote extracellular matrix production and that it may actively be involved in the pathogenesis of profibrotic disorders such as SSc.
Induction and production of these extracellular matrix proteins in response to hypoxia was time dependent and inversely correlated with oxygen levels [74]. Most of these proteins were significantly upregulated after 24 hours of oxygen deprivation, with a further significant increase after 48 hours. The expression of fibronectin-1, thrombospondin-1, COL1A2 and IGFBP-3 was significantly enhanced at 8% oxygen concentration and increased further with lower oxygen levels, reaching a maximum at 1% oxygen. Of note, severe and chronic hypoxia, as may be found in the skin of SSc patients [54], was associated with the most marked effects on the induction of extracellular matrix proteins.
These results were confirmed in a mouse model of systemic normobaric hypoxia [74]. Consistent with the results obtained in vitro, extracellular matrix proteins were upregulated in mice exposed to hypoxia after 24 hours compared with control mice breathing air with 21% oxygen. Prolonged exposure for 48 hours resulted in further upregulation of fibronectin 1, thrombospondin 1 and COL1A2, whereas TGF-βi and IGFBP3 mRNA levels decreased slightly. Because TGF-β is a major stimulus for the induction of extracellular matrix proteins in SSc [52,78], its role for hypoxia-dependent fibrogenesis was also studied in dermal SSc fibroblasts. Neutralizing antibodies against TGF-β completely abrogated the induction of COL1A2, fibronectin 1, thrombospondin 1 and TGF-βi in SSc fibroblasts that were cultured under hypoxic conditions for 48 hours [74]. These findings suggest that inhibition of TGF-β-dependent pathways may prevent the profibrotic effects of hypoxia.
Consistent with the results on TGF-β signalling, the expression of the fibrogenic cytokine connective tissue growth factor (CTGF) was also shown to be upregulated in SSc in response to hypoxia [79]. CTGF is a critical mediator of TGF-β-induced skin fibrosis in SSc [80]. Its serum levels are elevated in SSc patients and have been suggested to correlate with skin fibrosis [81]. Hong and coworkers [79] found increased levels of CTGF mRNA and protein in fibroblasts exposed to 1% of oxygen or treated with cobalt chloride, a chemical stabilizer of HIF-1α. The induction of CTGF in response to hypoxia depended on HIF-1α [79]. Because the authors concentrated on short-term hypoxia of up to 4 hours, it remains unclear whether CTGF is also induced by chronic hypoxia and by HIF-1α-independent mechanisms in SSc.
Thus, accumulating evidence suggests that hypoxia might be actively involved in pathogenesis of SSc by stimulating the release of extracellular matrix protein. This could result in a vicious circle of hypoxia and fibrosis. Hypoxia stimulates the production and accumulation of extracellular matrix. The resulting tissue fibrosis inhibits diffusion of oxygen, causing further tissue hypoxia, which stimulates further the production of extracellular matrix (Figure 1). Activation of TGF-βdependent pathways appears to play a central role in the induction of extracellular matrix proteins by hypoxia, and inhibition of TGF-β signalling might prevent hypoxia-induced tissue fibrosis. However, further studies are needed to characterize further the role played by hypoxia in SSc and to identify the molecular mechanisms activated by hypoxia in SSc.

Conclusions
Capillary rarification and disturbed blood flow, as well as excessive extracellular matrix accumulation, cause chronic tissue hypoxia in SSc. However, levels of HIF-1α protein are decreased, probably due to PHD-dependent negative feedback loops. Interestingly, physiological mechanisms to overcome tissue hypoxia are impaired and dysregulated in SSc. Insufficient angiogenesis and vasculogenesis cannot abolish tissue malnutrition and hypoxia. Compensatory overexpression of VEGF might even result in a futile vascular response to hypoxia, characterized by the chaotic vessel formation. Hypoxia stimulates the production of several extracellular matrix proteins in SSc fibroblasts in a time-and concentration-dependent manner. The excessive deposition of matrix might impair further the diffusion of oxygen and cause a vicious circle of hypoxia and tissue fibrosis. Currently, there are no specific modulators of HIFs or PHDs available for clinical use. Thus, it is not yet possible to target hypoxia selectively in SSc patients. However, because inhibition of TGF-β prevented the induction of extracellular matrix by hypoxia, blocking of TGF-β signalling might be one approach to target at least in part the hypoxia-induced matrix production in SSc.


extfieldsofstudy: Biology Medicine

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Mechanism for Amyloid Precursor-like Protein 2 Enhancement of Major Histocompatibility Complex Class I Molecule Degradation*

Earlier studies have demonstrated interaction of the murine major histocompatibility complex (MHC) class I molecule Kd with amyloid precursor-like protein 2 (APLP2), a ubiquitously expressed member of the amyloid precursor protein family. Our current findings indicate that APLP2 is internalized in a clathrin-dependent manner, as shown by utilization of inhibitors of the clathrin pathway. Furthermore, we demonstrated that APLP2 and Kd bind at the cell surface and are internalized together. The APLP2 cytoplasmic tail contains two overlapping consensus motifs for binding to the adaptor protein-2 complex, and mutation of a tyrosine shared by both motifs severely impaired APLP2 internalization and ability to promote Kd endocytosis. Upon increased expression of wild type APLP2, Kd molecules were predominantly directed to the lysosomes rather than recycled to the plasma membrane. These findings suggest a model in which APLP2 binds Kd at the plasma membrane, facilitates uptake of Kd in a clathrin-dependent manner, and routes the endocytosed Kd to the lysosomal degradation pathway. Thus, APLP2 has a multistep trafficking function that influences the expression of major histocompatibility complex class I molecules at the plasma membrane.

The efficacy of the cellular immune response to intracellular pathogens and tumors is reliant on major histocompatibility complex (MHC) 5 class I molecules. MHC class I molecules present peptides, including peptides derived from pathogens and tumors, at the cell surface to cytotoxic T lymphocytes.
Cytotoxic T lymphocytes have been selected during development for their ability to react to cells in the periphery expressing self MHC class I molecules bearing non-self peptides. The level of cell surface expression of MHC class I molecules on infected and malignant cells therefore dictates the extent to which the antigen-specific cytotoxic T lymphocytes can recognize, and subsequently lyse, abnormal cells. The cell surface expression of MHC class I molecules is controlled by the quantity and quality of MHC class I molecules that are assembled, and also by the rate at which MHC class I molecules depart from the plasma membrane.
Amyloid precursor-like protein 2 (APLP2) binds to the MHC class I molecule K d in cells that express ␤2-microglobulin (1,2). APLP2 regulates the level of folded K d molecules at the cell surface: a reduction in APLP2 expression causes an elevation in the amount of folded K d at the plasma membrane, and an increase in APLP2 expression results in a decline in folded K d surface expression (3,4). Higher APLP2 expression lowers the level of K d at the plasma membrane by increasing the endocytosis, destabilization, and turnover of K d molecules (4).
The family of proteins to which APLP2 belongs includes APL-1 in Caenorhabditis elegans, APPL in Drosophila, and three proteins in mammals: amyloid precursor protein (APP), amyloid precursor-like protein 1, and APLP2 (5,6). APLP2 shares a high degree of sequence homology with APP, particularly at the C-terminal end. However, APP possesses a unique ␤-amyloid peptide domain that is not present within APLP2 (6).
APLP2 is a ubiquitously expressed protein with both a type I transmembrane form and a secreted form generated by proteolytic cleavage (7,8). Functions for APLP2 have been shown in cell proliferation, cell adhesion, nerve growth, mitosis segregation, cell signaling, and transcriptional regulation (9 -14). The APLP2 signaling/transcriptional function is mediated by a C-terminal cleavage product, which translocates to the nucleus (14,15). By analogy to what is known from the study of the closely related protein APP, the cleavage of APLP2 that produces the free intracellular domain likely occurs after the arrival of the protein at the plasma membrane and may occur in endocytic compartments (16,17).
Our recent studies have demonstrated that, in addition to K d , APLP2 binds to other murine MHC class I molecules, with varied affinity (18). Furthermore, we have shown that APLP2 also binds to human MHC class I molecules, and that increased APLP2 expression causes down-regulation of human MHC class I molecules at the plasma membrane (19). The impact of APLP2 on cell surface expression of the MHC class I molecule, and the site of APLP2/MHC class I co-localization within the cell, depends on the specific MHC class I allotype (18). The polymorphic ␣1/␣2 domain and conserved (␣3/transmembrane/cytoplasmic) regions of the MHC class I molecule are involved in interaction with APLP2 (18).
Although our previous studies demonstrated that APLP2 is bound to the endocytosed K d molecule, the specific location where the binding occurs was not identified (4). In the present study, we established that APLP2 binds K d at the cell surface before these two proteins internalize into the same endocytic vesicles. Our results demonstrated that APLP2 is principally endocytosed by a clathrin-mediated process, because a dominant-negative dynamin II K44A mutant and the C-terminal portion of AP180, both of which inhibit clathrin-dependent endocytosis (20,21), blocked APLP2 internalization. Furthermore, clathrin-mediated endocytosis of APLP2 was required for APLP2 to promote K d endocytosis. The APLP2 cytoplasmic tail contains overlapping tyrosine-based NPXY and YXXØ motifs for adaptor protein-2 (AP-2) binding (22,23), and we demonstrated that mutation of the tyrosine included in both motifs impaired internalization of APLP2, indicating the importance of this sequence in APLP2 endocytosis. APLP2 mutated at this cytoplasmic domain tyrosine residue, in contrast to wild type APLP2, was unable to facilitate the endocytosis of K d . Cells expressing a high level of APLP2 were noted to have relatively few internalized K d molecules recycling to the plasma membrane, and delivery of K d to the lysosomes in these cells was shown by inhibition of lysosomal function with ammonium chloride. Our findings indicate that APLP2 is endocytosed by a clathrin-dependent mechanism, and that APLP2 utilizes clathrin-mediated endocytosis to increase the internalization of K d bound by APLP2 at the cell surface. Furthermore, our data demonstrate that APLP2 directs K d molecules to lysosomes and the K d molecules are degraded rather than recycled to the cell surface.

EXPERIMENTAL PROCEDURES
Antibodies (Abs)-Monoclonal Ab 34-1-2 recognizes the ␣1/␣2 domain of K d (24). The 64-3-7 monoclonal Ab can detect open, peptide-free L d (24,25) and can also detect open forms of other MHC class I heavy chains, such as K d , into which the 64-3-7 epitope has been introduced, and addition of the 64-3-7 epitope does not impair peptide presentation, trafficking, or surface expression of K d or other MHC class I molecules (26 -29). The 34-1-2 and 64-3-7 Abs were donated by Dr. T. Hansen (Washington University, St. Louis, MO). The Ab used for APLP2 detection was made against full-length APLP2 (Calbiochem). Rabbit anti-FLAG Ab was obtained from Cell Signaling, mouse anti-FLAG Ab was purchased from Sigma, and fluorescein isothiocyanate-conjugated goat anti-FLAG Ab was obtained from AbD Serotec. The rabbit anti-LAMP1 Ab was from Novus Biologicals. All Alexa Fluor secondary Abs used for immunofluorescence were purchased from Molecular Probes.
Cell Lines-Cells were cultured in RPMI 1640 medium (Invitrogen) containing glutamine, pyruvate, penicillin/streptomycin, and 15% fetal bovine serum. The HeLa cells were given by Dr. W. Maury (University of Iowa, Iowa City, IA). The HeLa cells were transfected with a cDNA for K d bearing the 64-3-7 epitope (etK d ) (26) in the pIRIS.puro2 vector (BD Biosciences Clontech) and were selected with puromycin. For some experiments, a C-terminal FLAG-tagged APLP2 cDNA in pCMVTag4A (Stratagene) was transiently transfected into HeLa-etK d to make HeLa-etK d cells that expressed APLP2 at higher levels. GFP-tagged Rab5 and Rab5Q79L cDNAs (30) in the EGFP vector (Clontech), kindly donated by Dr. R. Lodge, were transiently transfected into HeLa-etK d cells for some experiments. The GFP-dynamin II K44A mutant cDNA (20) in a pEGFP N1 vector (Clontech), used in transient transfections, was a gift from M. McNiven (Mayo Clinic, Rochester, MN). The FLAG-tagged C-terminal fragment of AP180 in the pFLAG-CMV2 vector that was used for transient transfections was contributed by Dr. J. Donaldson and Dr. L. Greene (National Institutes of Health, NICHD, Bethesda, MD). All transient transfections were done with Effectene (Qiagen), using 1 g of DNA per 0.5 ϫ 10 6 cells.
Immunoprecipitations and Western Blots-For immunoprecipitation and Western blotting, variations of published protocols (31) were used. For washes prior to lysis of cells, PBS containing 20 mM iodoacetamide (Sigma) was used, and the washes were done three times. Cells were lysed in CHAPS lysis buffer, which was 1% CHAPS (Roche Applied Science) in Tris-buffered saline (pH 7.4) with freshly added 0.2 mM phenylmethylsulfonyl fluoride and 20 mM iodoacetamide. For some experimental samples, the lysis buffer also contained a freshly added, saturating quantity of monoclonal Ab. For all samples, after 45 min with intermittent gentle mixing, the samples were centrifuged and the Protein A-Sepharose beads were washed in 0.1% CHAPS, 20 mM iodoacetamide in TBS (pH 7.4) four times and boiled in 0.125 M Tris (pH 6.8), 2% SDS, 12% glycerol, 0.02% bromphenol blue to elute the proteins. The eluates were electrophoresed on 4 -20% Tris glycine acrylamide gels (Invitrogen).
For Western blots, the proteins were transferred to Immobilon-P blotting membranes (Millipore). After blocking in reconstituted dry milk overnight at room temperature, the blots were incubated in a dilution of Ab for 2 h at room temperature, washed 3 times for 15 min with 0.05% Tween 20/PBS, and incubated 1 h in diluted biotin-conjugated goat anti-mouse or antirabbit IgG (Caltag Laboratories). Following 3 washes in 0.05% Tween 20/PBS, the blots were incubated 1 h in diluted streptavidin-conjugated horseradish peroxidase (Zymed Laboratories Inc.). The blots were washed 3 times for 15 min with 0.3% Tween 20/PBS and then treated with enhanced chemiluminescence Western blot reagents (GE Healthcare). The blots were exposed to Kodak Bio-Max film (Eastman Kodak Co., Rochester, NY) for visualization of the bands.
Immunoprecipitation and Western blotting were done to confirm the binding of the APLP2-Y755A-FLAG, APLP2-Y721A-FLAG, and APLP2-Y721A/Y755A-FLAG mutants to K d . For these experiments, the membranes were incubated in a dilution of Ab for 2 h at room temperature, washed 3 times with 0.05% Tween 20/PBS, and incubated for 1 h at room temperature in diluted horseradish peroxidase-conjugated goat antimouse Ab (Jackson ImmunoResearch). After three washes in 0.1% Tween 20/PBS, the membranes were treated with enhanced chemiluminescence reagents and exposed to film as described above.
Immunofluorescence Analysis-For immunofluorescence analysis, cells were grown on glass coverslips. For some experiments, cells were incubated at 37°C to allow internalization of proteins and Abs bound to them, and the remaining Abs still bound at the cell surface (not internalized) were removed by incubation in stripping buffer (0.5% acetic acid, 500 mM NaCl) for 90 s. The stripping step ensured that the subsequent immunofluorescence analysis would reveal only proteins that had been internalized. Cells were fixed with a solution of 4% (v/v) paraformaldehyde in PBS for 10 min. Ab incubations were done in staining solution (0.2% (w/v) saponin, 0.5% (w/v) bovine serum albumin/PBS) for 30 min at room temperature, unless otherwise specified. As a control for clathrin-mediated endocytosis, HeLa-etK d cells transfected with a cDNA for GFP-tagged dynamin II K44A or the FLAG-tagged AP180 C-terminal fragment were pulsed with fluorochrome-labeled transferrin (Tf-568 from Molecular Probes) for 5 min at 37°C.
In all immunofluorescence analysis experiments, PBS washes were 5 min at room temperature, and immediately prior to mounting for image analysis the cells were washed 3 times in PBS. All images were obtained with a Zeiss LSM 5 Pascal confocal microscope. A ϫ63 1.4 numerical aperture lens with appropriate filters was used. For some experiments, ImageJ software (rsb.info.nih.gov) was used to measure the fluorescence of the Ab-labeled internalized K d molecules. The mean fluorescence intensity was calculated with the histogram tool in ImageJ, using data from Ͼ80 cells of each type within a total of Ͼ10 images acquired with the same optical settings.

APLP2 and K d Bind at the Plasma Membrane and Are
Internalized Together-Previous studies from our laboratories showed that APLP2 co-localized in endosomes with K d internalized from the cell surface and that APLP2 could be co-immunoprecipitated with internalized K d (4). To determine whether the interaction between APLP2 and K d occurs at the cell surface, K d molecules at the surface of HeLa-etK d cells were labeled with the 34-1-2 Ab on ice for 20 min, and then the cells were lysed and Ab complexes isolated with Protein A-Sepharose. The complexes were electrophoresed, blotted, and probed with Abs to identify the K d heavy chain (Fig. 1A, top panel) or APLP2 (Fig. 1A, bottom panel). As controls, the same surface labeling procedure was performed using HeLa cells, and immunoprecipitations were done with Ab 34-1-2 using lysates of HeLa-etK d and HeLa cells that had not been surface labeled (Fig. 1A). From the lysates of HeLa-etK d cells labeled at the cell surface with 34-1-2 and maintained on ice to prevent internalization, APLP2 was co-immunoprecipitated with K d (Fig. 1A, bottom panel, lane 5). This approach specifically immunoprecipitated cell surface K d , as confirmed by the presence of only the high molecular weight (presumably glycosylated) K d in lane 5 (Fig. 1A, top panel).
As mentioned above, earlier studies from our laboratories showed that APLP2 and internalized K d co-localized in endosomes; however, these studies had not ascertained whether any of these APLP2 molecules were also internalized from the cell surface. To determine whether APLP2 co-localized with K d after both molecules were internalized from the cell surface, co-localization analysis was performed. HeLa-etK d cells were incubated with 34-1-2 and anti-APLP2 Abs for 12 min and the remaining Abs at the cell surface were removed. Cells were fixed, permeabilized, and stained with fluorescently labeled secondary Ab. As shown by the representative z-section in Fig.  1B, the K d and APLP2 molecules internalized from the cell surface were co-localized in the same vesicular compartments. To analyze whether the vesicles in which the internalized APLP2 and K d co-localized were endosomes, HeLa-etK d cells were transiently transfected with either GFP-Rab5 (Fig. 1C) or GFP-Rab5Q79L (Fig. 1D), and then surface labeled with Abs for K d and APLP2. Rab5 is an endosomal marker, and Rab5Q79L is a dominant-active Rab5 mutant that causes enlargement of early endosomes accessible to internalized proteins (30), thereby facilitating endosomal compartment identification by confocal microscopy. The internalization of Ab-labeled K d and APLP2 molecules was induced by incubating the cells at 37°C for 15 min. The remaining Ab at the cell surface was removed by acid washing, and the cells were permeabilized and stained with secondary Ab. APLP2 and K d , both endocytosed from the cell surface, were visualized within the same Rab5 ϩ endosomal vesicles.
Internalization of APLP2 by a Clathrin-dependent Mechanism and K d by a Clathrin-independent Mechanism-In an earlier study, we showed that increased expression of APLP2 reduces cell surface K d levels (4). As a step toward understanding the mechanisms by which elevated APLP2 expression increases the endocytosis of K d molecules, we assessed the means by which APLP2 itself is endocytosed. HeLa-etK d cells were transiently transfected with a dominant-negative dynamin II mutant, K44A, or with the C terminus of AP180. Both dynamin II K44A and the C-terminal portion of AP180 have been shown previously to inhibit clathrin-dependent endocytosis (20,21). To determine whether the dynamin II dominantnegative mutant or the C terminus of AP180 could down-regulate endocytosis of APLP2, each was transiently transfected into HeLa-etK d cells, and confocal microscopy was used to assess APLP2 internalization.
As shown in Fig. 2A, the expression of GFP-dynamin II K44A hindered the internalization of APLP2, indicating that APLP2 is endocytosed by a clathrin-dependent mechanism. Expression of GFP-dynamin II K44A also blocked the uptake of transferrin (included as a positive control), which is known to be a clathrinmediated process (data not shown). The presence of the C-terminal fragment of AP180-FLAG also prevented APLP2 and transferrin endocytosis ( Fig. 2B and data not shown), consistent with our results using the GFP-dynamin II mutant. (As a control, the level of surface staining of APLP2, without allowing APLP2 internalization, was assessed for comparison (supplemental Fig. S1).) In contrast, the endocytosis of K d was not inhibited by either GFP-dynamin II K44A or the C-terminal portion of AP180-FLAG (Fig. 2, C and D), suggesting K d is predominantly endocytosed in HeLa-etK d cells by a clathrin-independent pathway.
Internalization of APLP2 by a Clathrin-mediated Endocytic Mechanism Was Necessary for APLP2 to Facilitate K d Endocytosis-By binding to K d , APLP2 may recruit K d into the clathrin-mediated endocytic pathway as APLP2 itself is internalized. If so, then the tendency of APLP2 to increase K d endocytosis would depend upon the ability of APLP2 to be endocytosed. As a control, we incubated HeLa-K d cells that were transiently transfected with APLP2-FLAG with the 34-1-2 Ab to label the cell surface K d molecules and warmed the cells at 37°C to cause the K d molecules to be internalized. The remaining Abs at the surface were stripped off by an acid wash (with stripping buffer), and the cells were fixed and stained with anti-FLAG Ab. As expected from our previous study (4), cells that expressed APLP2-FLAG contained more endocytosed K d , relative to cells not expressing APLP2-FLAG (Fig. 3A). To determine whether APLP2 must be endocytosed for it to increase K d endocytosis, HeLa-etK d cells were transiently transfected with both APLP2-FLAG and the GFP-tagged dynamin II K44A dominant-negative mutant. The cells were pulsed with the 34-1-2 Ab to label the cell surface K d molecules and the K d molecules were allowed to internalize, then the remaining Abs at the surface were removed, and the cells were fixed and stained with anti-FLAG Ab. After washing, the cells were incubated with secondary Abs to identify the APLP2-FLAG and K d molecules (Fig. 3B). In contrast to the control results showing significantly higher numbers of K d molecules endocytosed in cells expressing APLP2-FLAG (in cells not transfected with GFP-dynamin II K44A) (Fig. 3C), the results displayed in Fig. 3D demonstrated that the expression of GFP-dynamin II K44A prevented enhancement of K d internalization by APLP2, as there was no difference in the quantity of K d molecules endocytosed in cells co-expressing both transfected APLP2-FLAG and GFP-dynamin II K44A, compared with cells expressing neither. These data suggest that overexpression of APLP2 causes enhanced endocytosis of K d through the clathrin pathway but expression of the dynamin II K44A mutant abolishes this effect. were identified as colocalized within vesicular compartments. Cell surface K d and APLP2 molecules were labeled with 34-1-2 Ab and anti-APLP2 Ab, respectively, for 12 min at 37°C to allow internalization of labeled proteins with the bound Abs, and any remaining Abs at the surface were removed. Cells were then fixed and stained with Alexa Fluor 568 goat anti-mouse Ab and Alexa Fluor 488 goat anti-rabbit Ab. Serial z-section images were acquired for HeLa-etK d cells, and the arrows point to common membrane structures on a representative z-section micrograph (obtained from six slices imaged at 0.4-m intervals). Red, K d ; green, ϭ APLP2; yellow, co-localized K d and APLP2. C and D, cell surface K d and APLP2 were internalized into the same endocytic vesicles. HeLa-etK d cells were transfected with either (C) GFP-Rab5 or (D) GFP-Rab5Q79L for 24 h, then pulsed with anti-K d Ab 34-1-2 and anti-APLP2 Ab and incubated in complete medium for 15 min at 37°C. Following the incubation, non-internalized Abs were removed. The cells were fixed, permeabilized, and stained with Alexa Fluor 568 goat anti-rabbit and Alexa Fluor 405 goat anti-mouse Abs. Red, APLP2; green, GFP-Rab5 or GFP-Rab5Q79L; blue, K d ; white, co-localized APLP2, K d , and GFP-Rab5 or GFP-Rab5Q79L. Bar, 10 m. The insets show a higher magnification of areas within the larger boxes, and the arrows point to representative endosomes with co-localized APLP2 and K d . DECEMBER 4, 2009 • VOLUME 284 • NUMBER 49

JOURNAL OF BIOLOGICAL CHEMISTRY 34299
A Tyrosine in the APLP2 C Terminus Was Required for the Effect of APLP2 on K d Internalization-NPXY and YXXØ sequence motifs present in other proteins bind to the AP-2 complex (21,22). These two endocytic motifs (NPXY and YXXØ) overlap within the amino acid sequence NPTYKYL of the APLP2 cytoplasmic tail. An additional YXXØ putative endocytic motif (in the sequence YGTI) is also present in the APLP2 cytoplasmic tail. We mutated the tyrosine residue in each of these sequences to an alanine (singly or together), and generated constructs with these APLP2 mutants (Y755A, Y721A, or Y721A/Y755A) bearing a C-terminal attached FLAG tag.
To verify that the tyrosine mutations did not disrupt binding of APLP2 to K d , HeLa-etK d cells were transiently transfected with vector only, wild type APLP2-FLAG, or each of the three mutant APLP2-FLAG constructs. The cells were lysed and immunoprecipitations were performed with anti-FLAG Ab to isolate FLAG-tagged proteins. The immunoprecipitates were Western blotted with anti-APLP2 Ab to confirm the quality of the immunoprecipitations or with 64-3-7 to detect the presence of co-immunoprecipitated K d . Results demonstrating K d binding to APLP2-Y755A-FLAG are shown in Fig. 4A, and similar results indicating K d association were obtained for APLP2-Y721A-FLAG and APLP2-Y721A/Y755A-FLAG (data not shown).
For assessment of the effect of the Y755A mutation on APLP2 endocytosis and on the ability of APLP2 to facilitate K d endocytosis, cells transfected with either wild type APLP2-FLAG or APLP2-Y755A-FLAG were incubated with anti-APLP2 Ab at 37°C to allow APLP2 internalization. Cell surface APLP2 was taken up normally in cells transfected with APLP2-FLAG (Fig. 4B, left panel). However, mutation of the Tyr-755 residue to alanine in APLP2 inhibited the internalization of cell surface APLP2 (Fig. 4B, right panel), indicating that Tyr-755 is required for APLP2 to internalize. In addition, it appears from these data that the expression of APLP2-Y755A-FLAG also likely inhibited the uptake of wild type, endogenous APLP2 (Fig. 4B, right panel). (Surface staining of APLP2, without warming the cells to permit internalization, was monitored as a control (supplemental Fig. S2).) We also found that the impairment of APLP2 endocytosis was very similar for APLP2-Y721A/Y755A-FLAG as compared with APLP2-Y755A-FLAG (data not shown). In contrast, we found no difference in APLP2 endocytosis between APLP2-Y721A- FIGURE 2. APLP2 was demonstrated to be endocytosed by a clathrin-mediated mechanism, but K d was shown to be endocytosed principally by a clathrin-independent mechanism. HeLa-etK d cells were transfected for 24 h with a cDNA encoding (A and C) GFP-tagged dynamin II K44A or (B and D) C-terminal fragment of AP180-FLAG. Cells were pulsed with (A and B) anti-APLP2 or (C and D) anti-K d Ab at 37°C for 20 min to allow Ab binding and internalization, and then the cells were stripped of non-internalized surface-bound Ab and fixed. Cells in B and D were stained with anti-FLAG Ab for 1 h in saponin-containing staining solution to visualize the AP180 C-terminal fragment (using rabbit anti-FLAG Ab if 34-1-2 Ab had been used for the pulse, or mouse anti-FLAG Ab if anti-APLP2 Ab had been used for the pulse). The cells transfected with GFP-dynamin II K44A were incubated with Alexa Fluor 568 goat anti-mouse secondary Ab to visualize internalized K d or with Alexa Fluor 568 goat anti-rabbit Ab to reveal internalized APLP2. For AP180-FLAG C terminus transfectants, the cells were incubated with Alexa Fluor 568 goat anti-mouse Ab and Alexa Fluor 488 goat anti-rabbit Ab if a 34-1-2 Ab pulse had been done, or with Alexa Fluor 488 goat anti-mouse Ab and Alexa Fluor 568 goat anti-rabbit Ab if an anti-APLP2 Ab pulse had been done. A and B, red, APLP2; C and D, red, K d ; A and C, green, GFP-dynamin II K44A; B and D, green, AP180 C terminus-FLAG. Bar, 10 m. Cells shown bordered with white are transfected with GFP-dynamin II K44A or AP180 C terminus-FLAG.
FLAG and wild type APLP2-FLAG (data not shown), indicating that the Tyr-721 position does not contribute to APLP2 internalization.
The APLP2 sequence NPTYKYL contains overlapping NPXY and YXXØ motifs, with Tyr-755 at the intersection of the two motifs. In studies on other proteins, it has been shown that both of these motifs can interact with AP subunits; however, only the NPXY motif has been demonstrated to interact with Dab2 (23, 32-34). We have found that trans-fection with AP-2 2 small interfering RNA largely abrogates internalization of APLP2, suggesting that APLP2 endocytosis is mediated by the AP-2 complex. In contrast, we found that Dab2 small interfering RNA transfection caused little decrease in APLP2 endocytosis, indicating that Dab2mediated clathrin-dependent endocytosis is a minor pathway for APLP2 internalization. However, transfection with both small interfering RNA for the AP-2 2 subunit and Dab2 resulted in greater inhibition of APLP2 endocytosis FIGURE 3. Blocking clathrin-mediated endocytosis by expression of dynamin II K44A prevented APLP2 from increasing K d endocytosis. A, HeLa-etK d cells transfected transiently with APLP2-FLAG (but not GFP-dynamin II K44A) for 24 h were pulsed with 34-1-2 Ab for 20 min in complete medium. At the end of the pulse period, non-internalized cell surface-bound Abs were removed. After fixation, the cells were stained with rabbit anti-FLAG Ab for 1 h. The cells were incubated with Alexa Fluor 568 goat anti-rabbit Ab to identify APLP2-FLAG and Alexa Fluor 405 goat anti-mouse Ab to identify 34-1-2-bound K d . Red, APLP2-FLAG; blue, K d . Bar corresponds to 10 m. Cells expressing APLP2-FLAG are encircled with a dashed white line. B, the same procedure described in A was followed except that the HeLa-etK d cells were co-transfected transiently with both APLP2-FLAG and GFP-dynamin II K44A for 24 h. Red, APLP2-FLAG; green, GFP-dynamin II K44A; blue, K d . Bar corresponds to 10 m. Cells co-expressing APLP2-FLAG and GFP-dynamin II K44A are encircled with a dashed white line. C, ImageJ software was used to measure the internalized K d fluorescence for Ͼ80 cells transfected with APLP2-FLAG versus no APLP2-FLAG. Note that in this control no cells were transfected with GFP-dynamin II K44A, and APLP2-FLAG was able to increase K d endocytosis. D, Image J software was used to measure the internalized K d fluorescence for Ͼ80 cells expressing both APLP2-FLAG and GFP-dynamin II K44A, as well as Ͼ80 cells expressing neither protein. In this case, some of the cells expressed GFP-dynamin II K44A, and APLP2-FLAG was not able to increase K d endocytosis if GFP-dynamin II K44A was also expressed in the same cell. C and D, mean fluorescence intensities and mean Ϯ S.E. were calculated, and p values were determined by Student's paired t test: C, p Ͻ 0.001; D, p ϭ 0.81. DECEMBER 4, 2009 • VOLUME 284 • NUMBER 49 than either small interfering RNA alone, evidently due to blockage of both major and minor pathways of APLP2 internalization (data not shown). Overall, these findings are con-sistent with a major role for the YXXØ motif and a more minor role for the NPXY motif in APLP2 endocytosis.

Effect of APLP2 on MHC Class I Endocytosis and Degradation
When HeLa-etK d cells expressing either APLP2-Y755A-FLAG or wild type APLP2-FLAG were labeled with anti-K d Ab and the cells were incubated at 37°C to induce K d endocytosis, the FLAG-tagged APLP2 Y755A, in contrast to wildtype APLP2-FLAG, did not promote K d endocytosis (Fig. 4, C and  D). A statistically significant difference was found between the number of endocytosed K d molecules in cells expressing APLP2-FLAG versus no APLP2-FLAG (Fig. 4E), but not between the number of internalized K d molecules in cells with APLP2-Y755A-FLAG versus no APLP2-Y755A-FLAG (Fig. 4F). Therefore, the Tyr-755-mediated endocytosis of APLP2 is necessary for APLP2 to facilitate K d endocytosis.
Elevated Expression of APLP2 Increased Shuttling of K d to the Lysosomes Rather Than Back to the Plasma Membrane-Our findings raised the question of what was the fate of the K d molecules that were shuttled to the clathrin-mediated endocytosis pathway by APLP2. For investigation of the subcellular itinerary of K d in cells that expressed lower and higher levels of APLP2, HeLa-etK d cells were transiently transfected with APLP2-FLAG, pulsed with anti-K d Ab 34-1-2 and warmed to 37°C, stripped of noninternalized Abs by acid wash, and either directly fixed (Fig. 5A) or first chased for 2 h and then fixed (Fig.  5B). Staining with anti-FLAG Ab and comparison of cells within the same fields that were not transfected versus transfected with APLP2 revealed that K d recycling after 2 h was decreased in cells expressing a higher level of APLP2 (Fig. 5B). In cells that were not transfected with APLP2-FLAG, by 2 h of chase most of the internalized 34-1-2-bound K d molecules had recycled back to the plasma membrane (indicated by arrows), although a few remained in vesicles within the cells (Fig. 5, B and C). However, in cells transfected with APLP2-FLAG (examples are shown encircled with a white Bar, 10 m. B, secondary Ab Alexa Fluor 488 (goat anti-mouse Ab) staining was done to identify APLP2-FLAGor APLP2-Y755A-FLAG-transfected cells, and the image shows the Alexa Fluor 568 (goat anti-rabbit) staining to visualize any internalized wild type APLP2-FLAG, APLP2-Y755A-FLAG, or wild type APLP2 (non-FLAG-tagged) molecules. C and D, Alexa Fluor 568 (goat anti-rabbit Ab) staining was used to identify APLP2-Y755A-FLAG or APLP2-FLAG, and Alexa Fluor 488 (goat anti-mouse Ab) was used to identify internalized, 34-1-2-bound K d . Red, APLP2-FLAG; green, K d ; yellow, co-localized APLP2-FLAG and K d . Cells encircled with white dashed lines are transfected with APLP2-FLAG (C) or APLP2-Y755A-FLAG (D). E, ImageJ software was used to measure the fluorescence of the internalized K d for Ͼ80 cells expressing APLP2-FLAG and Ͼ80 cells not expressing APLP2-FLAG. F, Image J software was used to measure the internalized K d fluorescence for Ͼ80 cells expressing APLP2-Y755A-FLAG and Ͼ80 cells not expressing APLP2-Y755A-FLAG. E and F, mean fluorescence intensities and mean Ϯ S.E. were calculated, and p values were determined by the Student's paired t test.
boundary), a majority of the 34-1-2-bound K d molecules were still localized to the cell interior at 2 h (Fig. 5, B and C). These data suggest that APLP2 not only enhances the endocytosis of K d but also down-regulates K d recycling.
Because K d recycling was decreased by APLP2, we next ascertained whether K d molecules were being transferred to the lysosomes. HeLa-etK d cells that had been transfected with APLP2-FLAG for 24 h were labeled with the anti-K d Ab 34-1-2 and incubated at 37°C. After K d uptake, non-internalized Abs were removed by acid wash, and the cells were subjected to a 4-h chase. The cells were then fixed and stained with anti-FLAG and anti-LAMP1 Abs for the transfected APLP2-FLAG and the LAMP1 lysosomal marker, respectively. Comparison of cells within the same fields that were transfected versus After the pulse period, non-internalized cell surface-bound Abs were removed. The cells were fixed, incubated for 1 h with rabbit anti-FLAG Ab, and stained with Alexa Fluor 488 goat anti-mouse Ab and Alexa Fluor 568 goat anti-rabbit Ab. B, the same procedure described in A was followed except that the cells were transferred to complete medium for 2 h after cell surface-bound Abs were removed with stripping buffer, and at the end of the 2-h chase period the cells were fixed and stained. Arrows indicate internalized K d that had recycled to the plasma membrane (A and B). Red, APLP2-FLAG; green, K d ; yellow, co-localized APLP2-FLAG and K d . Bar, 10 m. Cells outlined with white boundaries are transfected with APLP2-FLAG. C, comparison of intracellular K d in HeLa-etK d cells transfected with APLP2 versus untransfected cells in the same experiment. Quantification of mean fluorescence intensity of the stained intracellular K d was done with ImageJ software using data from Ͼ70 cells in each group. The graph shows the mean fluorescence intensity values, and the error bars on the graph indicate the mean Ϯ S.E. for each sample set. DECEMBER 4, 2009 • VOLUME 284 • NUMBER 49 untransfected with APLP2-FLAG revealed that the cells that had been transfected with APLP2-FLAG (marked with a boundary line) had few or none of the 34-1-2 Ab-labeled K d molecules still within the cells after the 4-h chase (Fig. 6A). With the cells that were not transfected with APLP2, most of the 34-1-2 Ab-labeled K d molecules had recycled back to the plasma membrane (shown by an arrow) by 4 h (Fig. 6A).

Effect of APLP2 on MHC Class I Endocytosis and Degradation
To confirm the destination of the K d molecules that were not being recycled to the plasma membrane, the cells were pretreated with the lysosome inhibitor ammonium chloride at 24 h post-transfection with APLP2-FLAG. Ab 34-1-2 was added to bind to the cell surface K d molecules, and the cells were warmed to cause K d endocytosis. Remaining non-internalized Abs were stripped off by an acid wash, and the cells were incubated for a 4-h chase before they were stained with anti-FLAG and anti-LAMP1 Abs. In ammonium chloride-treated cells that were not APLP2-FLAG-transfected, by 4 h most of the labeled K d molecules had recycled to the surface (Fig. 6B). Some intracellular vesicles in these non-APLP2-FLAG-transfected cells also contained labeled K d molecules that may have been rescued from degradation due to the inhibition of lysosomal function (Fig.  6B). In contrast, in cells transfected with APLP2-FLAG and treated with ammonium chloride, internalized K d molecules had accumulated in the lysosomes. This finding suggests that APLP2 had directed K d molecules to the lysosomes, but because lysosomal function was blocked by ammonium chloride, the K d molecules remained intact. Thus, in cells not treated with ammonium chloride, K d trafficking to lysosomes and (as a consequence) its turnover in the lysosomes was increased upon elevation of the cellular level of APLP2.

DISCUSSION
Endocytosis of MHC class I molecules has been studied using several cell types as models, including lymphocytes, fibroblasts, and macrophages, with variability reported for the extent and pathway of internalization (35)(36)(37)(38)(39)(40)(41)(42)(43)(44)(45)(46). The endocytosis of MHC class I molecules is dependent on a sequence in the MHC class I molecule cytoplasmic tail (40,41,47). Studies have indicated that MHC class I molecules are recycled to the cell surface after internalization (e.g. Ref. 42) by a process in which Eps15 homology domain-containing proteins (EHD1 and EHD4) play an important role (48 -50). EHD1 causes the generation of tubules that contain endocytosed MHC class I molecules, and higher expression of EHD1 increases MHC class I recycling, whereas reduced EHD1 expression slows the rate of MHC class I recycling (48,49).
In our studies, the endocytosis of K d in HeLa-etK d cells was not inhibited by dynamin II K44A or the C-terminal portion of AP180, indicating that endocytosis was primarily non-clathrin mediated. This observation with K d is in agreement with previously published findings that the endocytosis of other MHC class I molecules occurs principally via a clathrin-independent path (35,44), although there have also been reports that MHC class I molecules are internalized by a clathrin-dependent pathway in certain cell types (38,39). Our observation that APLP2 can divert K d into the clathrin-dependent pathway suggests FIGURE 6. K d routing to lysosomes for degradation was increased in cells with a higher level of APLP2. A, HeLa-etK d cells transfected with APLP2-FLAG for 24 h were pulsed for 15 min at 37°C with anti-K d Ab 34-1-2. After the pulse, non-internalized cell surface-bound Abs were removed. The cells were transferred to complete medium for 4 h, and then fixed and stained with fluorescein isothiocyanate-conjugated goat anti-FLAG Ab and rabbit anti-LAMP1 Ab for 1 h. The cells were stained with Alexa Fluor 405 goat anti-mouse Ab (to identify the 34-1-2-labeled K d molecules) and Alexa Fluor 568 goat anti-rabbit Ab (to identify the LAMP1 marker in the lysosomes). Green, APLP2-FLAG; blue, K d ; red, LAMP-1; white, co-localized APLP2-FLAG, K d , and LAMP-1. Bar, 10 m. B, the same procedure as in A was followed except that at 24 h the cells were treated with 20 mM ammonium chloride for 12 h before the 34-1-2 Ab pulse. The insets show a higher magnification of areas within the larger boxes, and the arrows point to representative lysosomes with co-localized APLP2-FLAG and K d .
that the intervention of APLP2 or other non-MHC proteins in some cells may explain some of the differences in the pathways of MHC class I internalization that have been described.
Our findings showed that APLP2 endocytosis is dependent on the tyrosine at position 755, which is located within overlapping NPXY and YXXØ motifs in the distal part of the APLP2 cytoplasmic domain. The Tyr-755 and both motifs are conserved between mouse and human APLP2. Furthermore, the signal for endocytosis of the closely related molecule APP has also been shown to rely on motifs in the cytoplasmic tail. The C-terminal domain of APP contains two overlapping NPXY and YXXØ endocytic motifs within the sequence YENPTYKFF, and deletion of the YENPTY sequence in APP decreased APP endocytosis (51). Consistent with this finding, studies from several laboratories have shown that cells expressing either APP with the cytoplasmic domain deleted or APP mutated at the tyrosine in the NPTY cytoplasmic tail sequence secrete higher levels of soluble APP than cells expressing wild type APP, which could result from impairment of APP endocytosis (52)(53)(54). Subsequent mutational analysis defined the YENP sequence as the critical tetrapeptide motif for APP internalization and the tyrosine in the NPTY sequence as a lysosomal targeting signal for APP (55). Besides the dominant YENPTY sequence, the cytoplasmic tail of APP contains a YTSI sequence proximal to the transmembrane region that has another tyrosine-based endocytic motif (YXXI). Mutation of the tyrosine in the YTSI sequence does not impair APP endocytosis, although this motif plays an important role in the basolateral targeting of APP in polarized cells (56).
In a previous study from our laboratories (4), APLP2 was discovered to interact with folded, internalized K d molecules and facilitate K d endocytosis. A study focused mainly on the closely related protein APP has been recently published (57), in which both APLP2 and APP were observed to interact with the high-affinity choline transporter, and APP was demonstrated to facilitate its endocytosis. Together, our findings and those of Wang et al. (57) suggest that APP family members APP and APLP2 play important roles in regulating expression of cell surface proteins. In our model of APLP2 regulation of K d surface expression, APLP2 binds K d at the plasma membrane (Fig. 7, A and B). Via the clathrin-mediated and AP-2-depen- FIGURE 7. Diagram of a model in which increased expression of APLP2 results in the recruitment of clathrin machinery at APLP2⅐K d complexes, causing uptake of K d in a clathrin-dependent manner and routing K d to the lysosomes. A, increased availability of APLP2 leads to more binding of APLP2 to K d and subsequent internalization of both APLP2 and K d by a clathrin-mediated mechanism into Rab5 ϩ early endosomes. B, inhibition of APLP2 clathrin-mediated endocytosis by mutation of Tyr-755 to alanine (as depicted) or by dynamin II K44A or the AP180 C-terminal fragment (not depicted in this figure) prevents APLP2 from enhancing K d endocytosis. C, in cells with less APLP2, the majority of K d molecules follow a recycling pathway from the Rab5 ϩ endosomes to the endocytic recycling compartment (ERC) and back to the cell surface (indicated by the thick arrows) and a minority traffic to late endosomes (LE) and lysosomes (LYS) (indicated by the thin arrows). D, in cells with a higher level of APLP2, the majority of K d molecules follow a degradation pathway from the Rab5 ϩ endosomes to the lysosomes (LYS) (indicated by thick arrows), and a minority of K d molecules enter a recycling pathway (indicated by thin arrows). dent endocytosis (involving the Y755 in the APLP2 C terminus) of APLP2, APLP2 increases the internalization of K d , diverting it away from its clathrin-independent route of endocytosis (Fig.  7, A and B). APLP2 increases the trafficking of the endocytosed K d molecules to the late endocytic compartment and lysosomes, where the K d molecules are degraded (Fig. 7, C and D). Thus, despite convergence at Rab5 ϩ early endosomes, the many K d molecules escorted into the cell interior by APLP2 molecules as they take the clathrin pathway are principally destined for lysosomal destruction, whereas the fewer K d molecules internalized via the clathrin-independent pathway when a lower level of APLP2 is expressed in the cell are predominantly recycled (Fig. 7, C and D).
APLP2 has been reported to be overexpressed in tumors (58), and APLP2 levels were noted to be higher in invasive breast cancer lesions than in non-invasive breast cancer lesions (59). Therefore, the effects of increased APLP2 on MHC class I molecules that we have demonstrated have possible implications not only in normal MHC class I molecule turnover but also in cancer evasion of the cellular immune response. Recent findings from our laboratory indicating that APLP2 is highly expressed in many human tumor cell lines and decreases the cell surface expression of human MHC class I molecules on tumor cells are consistent with this supposition (19). In addition, other functions of APLP2 related to cell signaling, adhesion, migration, and proliferation, which may play a role both in normal cell physiology and in malignancy, may all be influenced, directly or indirectly, by the endocytosis of APLP2. Overall, our analyses of APLP2-directed endocytosis of MHC class I molecules provide significant new perspectives on crucial cellular processes.