Mice lacking asparaginyl endopeptidase develop disorders resembling hemophagocytic syndrome

Mice lacking asparaginyl endopeptidase develop disorders resembling hemophagocytic syndrome Chi-Bun Chana,1, Michiyo Abeb,1, Noriyoshi Hashimotob, Chunhai Haoa, Ifor R. Williamsa, Xia Liua, Shinji Nakaoc, Akitsugu Yamamotod, Chengyun Zhenge,f, Jan-Inge Hentere, Marie Meethse,f, Magnus Nordenskjoldf, Shi-Yong Lia, Ikuko Hara-Nishimurag, Masahide Asanob,2, and Keqiang Yea,2 aDepartment of Pathology and Laboratory Medicine, Emory University School of Medicine, 615 Michael Street, Atlanta, GA 30322; bDivision of Transgenic Animal Science, Advanced Science Research Center, and cDepartment of Cellular Transplantation Biology, Graduate School of Medical Science, Kanazawa University, 13-1 Takara-machi, Kanazawa 920-8640, Japan; dDepartment of Bio-Science, Nagahama Institute of Bio-Science and Technology, 1266 Tamura-machi, Nagahama 526-0829, Japan; gDepartment of Botany, Graduate School of Science, Kyoto University, Kitashirakawa, Sakyo-ku, Kyoto 606-8502, Japan; eDepartment of Woman and Child Health, Childhood Cancer Research Unit, and fDepartment of Molecular Medicine and Surgery, Clinical Genetics Unit, Karolinska Institute, Karolinska University Hospital, SE-141 86 Stockholm, Sweden

Edited by Solomon H. Snyder, Johns Hopkins University School of Medicine, Baltimore, MD, and approved November 10, 2008 (received for review October 1, 2008)

Asparaginyl endopeptidase (AEP or legumain) is a lysosomal cysteine protease that cleaves protein substrates on the C-terminal side of asparagine. AEP plays a pivotal role in the endosome/ lysosomal degradation system and is implicated in antigen processing. The processing of the lysosomal proteases cathepsins in kidney is completely defective in AEP-deficient mice with accumulation of macromolecules in the lysosomes, which is typically seen in lysosomal disorders. Here we show that mutant mice lacking AEP develop fever, cytopenia, hepatosplenomegaly, and hemophagocytosis, which are primary pathological manifestations of hemophagocytic syndrome/hemophagocytic lymphohistiocytosis (HLH). Moreover, AEP deficiency provokes extramedullary hematopoiesis in the spleen and abnormally enlarged histiocytes with ingested red blood cells (RBCs) in bone marrow. Interestingly, RBCs from AEP-null mice are defective in plasma membrane components. Further, AEP-null mice display lower natural killer cell activity, but none of the major cytokines is substantially abnormal. These results indicate that AEP might be a previously unrecognized component in HLH pathophysiology. hematopoiesis 兩 macrophage 兩 legumain 兩 lysosomal disorder

H

emophagocytic syndrome (hemophagocytic lymphohistiocytosis, HLH) is a life-threatening condition caused by hyperinflammatory response. It can be classified into familial and acquired forms (1). The primary symptoms of HLH include prolonged fever, cytopenia, hepatosplenomegaly, and hemophagocytosis by activated macrophages (2). Whereas acquired HLH is induced by a variety of infectious organisms, such as viruses, bacteria, protozoa, or fungi (1), genetic HLH is a disorder inherited in an autosomal recessive or X-linked manner. Several loci have been implicated in the pathophysiology of genetic HLH, including PRF1 (3), UNC13D (4), RAB27A (5), LYST (6), and SH2D1A (7, 8). Most of these known defective genes associated with HLH are involved in the cytotoxic-granule exocytosis pathway of T lymphocytes, suggesting the critical role of the lysosomal granule in maintaining normal cellular immunity. For example, mutation of UNC13D in human impairs the release of granzyme and perforin from cytolytic granules at the immunological synapse (4). Nevertheless, up to 70% of familial HLH is caused by unknown genetic defects. The identification of new genes that are involved in the development of HLH would thus be beneficial to our understanding of the disease pathophysiology and our current therapeutic regimen to treat HLH. Asparagyl endopeptidase (AEP or legumain) is a lysosomal cysteine protease that cleaves protein substrates on the Cterminal side of asparagine (7, 8). AEP is highly expressed in kidney and localizes in the late endosomes and lysosomes of the kidney-proximal tubule cells. Disruption of AEP leads to late endosomes and lysosomes augmentation and dislocation from

468 – 473 兩 PNAS 兩 January 13, 2009 兩 vol. 106 兩 no. 2

the apical region of the kidney-proximal tubule cells and the abnormal lysosomes contained in electron-dense and/or membranous materials (9, 10). AEP activation is autocatalytic and requires sequential removal of C- and N-terminal propeptides at different pH thresholds. Recently, we have reported that neuronal AEP is involved in neuronal apoptosis by degrading the DNase inhibitor SET during excitoneurotoxicity (11). AEP also participates in antigen presentation, the activity of which is inversely proportional to myelin basic protein (MBP) epitope presentation (9, 10). In this report, we show that AEP-knockout mice develop classical syndromes resembling HLH with extramedullary hematopoiesis in spleen and abnormally enlarged histiocytes with ingested RBCs in bone marrow. Moreover, higher body temperature and reduced natural killer cell activity are observed in AEP-null mice. These results indicate that AEP might be a previously unrecognized component in HLH pathology. Results AEP-null mice are fertile and viable with no overt behavioral abnormality, although their body weights are reduced compared with wild-type littermates (9, 10). We observed a significant enhanced body temperature in AEP-null mice compared with their control littermates (Fig. 1A). To clarify the cause of fever, we determined various inflammatory cytokine levels in the serum. TNF-␣ levels were increased in some of AEP⫺/⫺ mice, suggesting that the fever is in part caused by increased TNF-␣ levels. However, most of AEP⫺/⫺ mice showed normal inflammatory cytokine levels (Fig. S1). A complete blood count revealed that AEP-null mice, but not the control littermates, had marked low concentration of RBCs and hemoglobin, suggestive of severe anemia (Table 1). We also compared the peripheral blood parameters from different ages of AEP ⫹/⫺ and ⫺/⫺ mice and found that the hematocrit values from AEP-null mice were gradually decreased in an age-dependent manner (Fig. 1B). On the other hand, the reticulocyte percentage progressively inAuthor contributions: C.-B.C., M. Abe, N.H., C.H., I.R.W., C.Z., J.-I.H., M. Asano, and K.Y. designed research; C.-B.C., M. Abe, N.H., C.H., I.R.W., X.L., S.N., A.Y., and M.M. performed research; I.H.-N. contributed new reagents/analytic tools; C.-B.C., M. Abe, N.H., C.H., I.R.W., C.Z., M.M., M.N., S.-Y.L., M. Asano, and K.Y. analyzed data; and C.-B.C., M. Asano, and K.Y. wrote the paper. The authors declare no conflict of interest. This article is a PNAS Direct Submission. 1C.-B.C.

and M. Abe contributed equally to this work.

2To

whom correspondence may be addressed. E-mail: [email protected] or [email protected].

This article contains supporting information online at www.pnas.org/cgi/content/full/ 0809824105/DCSupplemental. © 2008 by The National Academy of Sciences of the USA

www.pnas.org兾cgi兾doi兾10.1073兾pnas.0809824105

B

38 37 36

40 35 30 25 20

+/-

-/-

3

6 15 Age (month)

D 350

20

+/+ -/-

300 15 EPO (ng/ml)

Ratio of reticulocytes (%)

*

45 ***

35

C

Table 1. Complete blood count of AEP-deficient mice at the age of 6 months

50

** *

10 5

250

* *

3

6 15 Age (month) +/+

150

⫺/⫺ (n ⫽ 8)

Body weight, g Spleen weight, mg RBCs, 104/␮L Hemoglobin, g/dL Hematocrit, % Mean corpuscular volume, fL Mean corpuscular hemoglobin, pg Mean corpuscular hemoglobin conc., g/dL Total WBCs, 102/␮L Lymphocytes, 102/␮L Monocytes, 102/␮L Neutrophils, 102/␮L Platelets, 104/␮L

30.9 ⫾ 3.1 153 ⫾ 45 971 ⫾ 50 13.8 ⫾ 0.6 47.2 ⫾ 1.8 48.6 ⫾ 1.2 14.2 ⫾ 0.4 29.2 ⫾ 0.4 36.5 ⫾ 13.8 23.6 ⫾ 13.2 6.0 ⫾ 2.2 6.6 ⫾ 5.3 79.8 ⫾ 16.5

28.2 ⫾ 3.7 330 ⫾ 142† 798 ⫾ 120‡ 10.9 ⫾ 1.6‡ 38.2 ⫾ 5.1‡ 48.0 ⫾ 1.0 13.7 ⫾ 0.3* 28.7 ⫾ 0.5* 36.1 ⫾ 9.7 21.3 ⫾ 5.6 9.1 ⫾ 5.3 5.5 ⫾ 1.8 58.2 ⫾ 33.4

100 0

2

4 8 Age (month) -/-

E

IHC: anti-EPOR Fig. 1. Higher body temperature and anemia in AEP-deficient mice. (A) Body temperature of AEP-knockout (filled circles) and control heterozygous (open circles) mice at 12 months of age. ⴱⴱⴱ, P ⬍ 0.005, Student’s t test, ⫹/⫺, n ⫽ 4, ⫺/⫺, n ⫽ 7. Body core temperature was continuously recorded on a potentiometer by a copper– constantan thermocouple covered with polyethylene tubing (outside diameter 1 mm). (B) Age-dependent decrease of circulating hematocrit was seen in AEP-null (filled circles) animals but not in the heterozygous control mice (open circles). ⴱ, P ⬍ 0.05, Student’s t test, 3-monthold: ⫹/⫺, n ⫽ 6, ⫺/⫺, n ⫽ 6; 6-month-old: ⫹/⫺, n ⫽ 7, ⫺/⫺, n ⫽ 6; 15-month-old: ⫹/⫺, n ⫽ 2, ⫺/⫺, n ⫽ 4. (C) Age-dependent increase of circulating reticulocytes was seen in AEP-null (filled circles) animals but not in the heterozygous control mice (open circles) ⴱ, P ⬍ 0.05; ⴱⴱ, P ⬍ 0.01, Student’s t test). Data were obtained from the same mice examined in B. (D) Circulating Epo from wildtype (filled bars) and AEP-knockout (open bars) were determined by ELISA. A progressive increase of serum Epo was recorded in AEP-null animals with increased age. ⴱ, P ⬍ 0.05, Student’s t test, n ⫽ 6. (E) Bone marrow sections from femur of wild-type (⫹/⫹) and AEP-knockout (⫺/⫺) animals were stained with EpoR-specific antibody. IHC, immunohistochemistry. Enhanced EpoR expression was detected in AEP-knockout mice. (Scale bar, 20 ␮m.)

creased in AEP⫺/⫺ mice compared with control mice, fitting with the observation of anemia in AEP-deficient mice (Fig. 1C). In contrast, the total number of white blood cells (WBCs) increases as the AEP-knockout mice get older, which is caused predominantly by leukocytosis (Table 2). Erythropoietin (Epo), which is produced by the kidney in the adult and by the liver in the fetus, increases RBCs through supporting the survival of erythroid progenitor cells and stimulating their differentiation and proliferation via binding to Epo receptors (EpoRs) present on the surface of immature erythroid cells (12). Defects in Epo production lead to severe anemia caused by the absence of circulating RBCs. To investigate whether anemia in AEPChan et al.

⫹/⫺ (n ⫽ 8)

Data are expressed as mean ⫾ SD. *, P ⬍ 0.05; †, P ⬍ 0.01; ‡, P ⬍ 0.005; Student’s t test, n ⫽ 8.

200

50 0

Parameter

deficient mice is caused by a lack of Epo production, we determined the concentrations of Epo in the blood. Surprisingly, Epo production was markedly increased in an age-dependent manner in both types of mice, but the circulating Epo was significantly higher in AEP-lacking mice than in wild-type mice (Fig. 1D). Concomitantly, EpoR was dramatically increased in bone marrow from AEP⫺/⫺ mice compared with wild-type mice, fitting with the Epo increase in blood (Fig. 1E). Presumably, the elevated Epo and EpoR might compensate for the RBC loss in the AEP-lacking mice. Next, we monitored the weight of different organs from the age-matched wild-type and knockout mice and found that the spleen was substantially larger in AEP⫺/⫺ mice than in control mice. The liver was also significantly enlarged in AEP⫺/⫺ mice as compared with the normal counterparts, whereas other organs were comparable in size and weight. On average, the spleens from AEP-null mice were 5–10 times larger than those of wild-type controls (Fig. 2A). The difference in organ size was age dependent. As AEP⫺/⫺ mice grew older, the spleen grew much bigger than that of the control mice (Fig. 2B). Noticeably, the color of the spleen and liver from AEP-null mice was much darker than the wild-type control (Fig. S2), suggestive of an increase in the RBCs in the organs. To test this possibility, we examined the histology of the spleen and found a significant increase in extramedullary hematopoiesis in the AEP⫺/⫺ spleen as evident in an increase of morphologically normal megakaryocytes (Fig. 2C). Moreover, a substantial number of erythrocytes (Fig. 2D) and immature myeloid lineage cells (Fig. 2E) were observed in the AEP⫺/⫺ spleen, further supporting that extramedullary hematopoiesis was in progress. These findings indicate the extramedullary hematopoiesis might be the cause of splenomegaly. However, the liver histology appeared normal (data not shown). To confirm the hematopoiesis, the spleen cells from AEP-null mice were subjected to hypotonic lysis of mature erythrocytes Table 2. WBC contents of AEP-deficient mice at the age of 12 months Parameter 102/␮L

Total WBCs, Lymphocytes, 102/␮L Monocytes, 102/␮L Neutrophils, 102/␮L

⫹/⫺ (n ⫽ 5)

⫺/⫺ (n ⫽ 5)

40.0 ⫾ 11.4 20.6 ⫾ 8.3 4.4 ⫾ 2.9 15.1 ⫾ 6.9

115 ⫾ 36* 69.4 ⫾ 26.4* 20.3 ⫾ 20.8 25.0 ⫾ 19.5

Data are expressed as mean ⫾ SD. *, P ⬍ 0.05, Student’s t test, n ⫽ 5. PNAS 兩 January 13, 2009 兩 vol. 106 兩 no. 2 兩 469

CELL BIOLOGY

39

Hematocrit (%)

Body Temperature (ºC)

A

Pancreas

Lung

Kidney

Stomach

Liver

Spleen

***

B 2.0 Spleen weight (g)

+/+ -/-

*

Heart

8 7 6 5 4 3 2 1 0

Brain

% Body weight

A

+/-/-

1.5 1.0 0.5 0.0 0

3

6

9 12 Age (month)

15

18

D

+/-

C

-/-

E

H&E staining Fig. 2. Hepatosplenomegaly in AEP-knockout mice. (A) The weights of both the liver and spleen from AEP-null animals (open bar) were significantly higher than those of age-matched wild-type controls (filled bar). ⴱ, P ⬍ 0.05; ⴱⴱⴱ, P ⬍ 0.001, n ⫽ 8. (B) Age-dependent increase of spleen weight in AEP-knockout mice. AEPknockout mice (filled circles) and heterozygous control (open circles) of various ages as indicated were killed and the spleens were weighed (3-month-old: ⫹/⫺, n ⫽ 6, ⫺/⫺, n ⫽ 6; 6-month-old: ⫹/⫺, n ⫽ 6, ⫺/⫺, n ⫽ 6; 9-month-old: ⫹/⫺, n ⫽ 3, ⫺/⫺, n ⫽ 10; 12-month-old: ⫹/⫺, n ⫽ 4, ⫺/⫺, n ⫽ 3; 15-month-old: ⫹/⫺, n ⫽ 3, ⫺/⫺, n ⫽ 10; 18-month-old: ⫹/⫺, n ⫽ 6, ⫺/⫺, n ⫽ 8.). (C) H&E staining of the spleen from both AEP-heterozygous (⫹/⫺) and homozygous mice (⫺/⫺). (Scale bar, 500 ␮m.) (D) H&E staining of AEP-null spleen showing numerous hemophagocytes. (Scale bar, 500 ␮m.) (E) Giemsa staining of splenocytes from AEP⫺/⫺ mice showing immature myeloid lineage cells.

and analyzed by flow cytometric immunophenotyping for nucleated hematopoiesis precursor cells. As the weight of the spleen from AEP-null mice increased, the ratio of CD3 and B220 double-negative cells was markedly elevated (Fig. 3A), indicating that non-T cell and non-B cell populations were augmented as splenomegaly was getting worse. To identify the non-T cell and non-B cell populations, FACS analysis was further performed. The percentage of Ter-119⫹/CD45⫹ erythroid lineage cells in the spleen was dramatically increased in the AEP-null mice (e.g., 18.2% in a representative AEP-null mouse compared with just 0.4% in a wild-type control) (Fig. 3B). Myeloid cells identified by Gr-1 and Mac-1 staining were also increased in the AEP-null mice (e.g., 10.2% in AEP-null mice versus 1.2% in wild-type) (Fig. 3C). The presence of a large number of erythroid lineage Ter-119⫹ cells in the spleens of AEP-null mice confirms that extramedullary hematopoiesis was taking place. Compared with the wild-type mice, the bone marrow in AEP-null mice was infiltrated with many enlarged histiocytes that displayed a small and eccentrically placed nucleus and a cytoplasm with characteristic crinkles or striations, resembling Gaucher cells in Gaucher disease, a congenital storage disease due to lysosomal enzyme defect. We made the same observation in the spleen of AEP-null mice (Fig. 4A). Numerous histiocytes revealed ingested RBCs on one side and striated cytoplasm on the other side, suggesting accumulation of lipids from the digested RBCs. Hemophagocytes were observed in the bone marrow of AEP⫺/⫺ mice 470 兩 www.pnas.org兾cgi兾doi兾10.1073兾pnas.0809824105

at 3 months of age and they increased remarkably with age (Fig. S3A). These Gaucher-like cells showed strong autofluorescence as well as engulfed erythrocytes (Fig. S3D). Periodic acid/Schiff reagent (PAS) staining further supported that these were Gaucherlike cells (Fig. S3E). Hemophagocytes observed in the bone marrow of AEP⫺/⫺ mice were further examined by Cytospin. These hemophagocytes were both F4/80 and CD68-positive histiocytes (Fig. S3F), confirming their identities as macrophages. Electron microscopic examination of bone marrow showed that the aberrant histiocytes engorged numerous RBCs, many of which were apparently intact, but some of which were partially or completely degraded (Fig. 4B). Fluorescent microsphere phagocytic assay with the peritoneal macrophages demonstrated that AEP depletion evidently enhanced the phagocytic activity, suggesting that phagocytic activity of macrophages (histiocytes) in AEP⫺/⫺ mice was elevated (Fig. 4C). To further explore whether the hemophagocytic activity in AEP⫺/⫺ mice results from augmented RBC engulfing activity by macrophages, we isolated RBCs and peritoneal macrophages from AEP⫺/⫺ and ⫹/⫺ mice. At 2 months of age, the macrophages from both mice engulfed opsonized RBCs with comparable activities (data not shown). By contrast, at 12 months of age, the macrophages from AEP⫺/⫺ mice engulfed more opsonized RBCs, but not unopsonized RBCs, than those from AEP⫹/⫺ mice, no matter whether the RBCs were isolated from AEP⫺/⫺ or ⫹/⫺ mice, confirming that phagocytic activity of macrophages from AEP⫺/⫺ mice was elevated (Fig. 4D). To examine whether unopsonized RBCs are engulfed by macrophages, macrophages and RBCs were incubated for 24 h instead of 1 h. Phagocytic activity of macrophages from AEP⫺/⫺ mice against unopsonized RBCs was elevated compared with macrophages from AEP⫹/⫺ mice, regardless of the genotype of RBCs, although the phagocytic index was very low (Fig. S4A). In contrast to the enhanced macrophage activity, the natural killer cell activity in the AEP-null animal is significantly reduced (Fig. 4E). The histological observation that the histiocytes specifically engulfed mature RBCs suggests the possible defect in the RBCs. To explore whether the plasma membrane in RBCs from AEP⫺/⫺ mice contains any defect, we monitored the resolving pattern of RBC membrane proteins on SDS/PAGE and found that the some of the erythrocyte proteins in AEP-null mice displayed an age-dependent reduction compared with those from wild-type mice (Fig. S4B). Proteomic analysis of the proteins from AEP-null circulating erythrocytes demonstrated that valosin-containing protein, MPP1, and ␣-subunit of ATP synthase were substantially altered in AEP-null mice. We further performed immunoblotting analysis to confirm the proteomic findings. As a control, the RBC membrane-associated protein 4.1R remained stable in both wild-type and AEP-knockout mice. In contrast, EpoR was substantially decreased in AEP-null mice and valosin-containing protein was completely lost in AEPdepleted mice, whereas they remained stable in the wild-type control (Fig. S4C). The deficiency of EpoR in RBCs from AEP-null mice at least in part explains why AEP-lacking mice develop anemia. The loss of EpoR and valosin-containing protein in AEP-null erythrocyte membrane might result in the deformity of the RBCs and phagocytosis by histiocytes once they are produced in bone marrow. Discussion Our results show that AEP-knockout mice develop major characteristic features of HLH. Clinically, AEP-null animals have prolonged body temperature elevation and progressive hepatosplenomegaly. Histologically, hemophagocytosis in both the bone marrow and spleen were seen in AEP-knockout mice. Moreover, AEP-knockout mice display cytopenia-reduced NK cell activity. These hallmarks for HLH suggest that AEP might play an unexplored role in development of HLH. Chan et al.

A

+/-: 0.075g

-/-: 0.332g

-/-: 0.904g

42.71

46.96 41.31

35.22

4.13

32.49 20.98

60.02

81.06

7.21

14.65

B 220

9.84

-/-: 4.107g

C D3

B

-/18.2 0.1

0.4

33.7

98.2

-/-

+/+ 10.2

1.2

Gr-1

Ter119

36.9

C

+/+

CD45

77.8

11.8

95.6

3.0

Mac-1

Although the phenotypes displayed in AEP-null mice highly resemble the primary clinical symptoms associated with HLH, no significant elevation of circulating cytokine is detected in AEP-knockout mice as shown in HLH patients (13, 14). A possible explanation to this discrepancy lies in the timing of the disease onset. HLH can be a rapid fatal condition, but the symptoms may present with less severe signs. Since AEPknockout mice show no aberrant death in early age (data not shown), it is thus suggested that the HLH development in AEP-null mice progresses slowly. The serum cytokine levels are not statistically changed in AEP-null mice up to 18 months of age. Nevertheless, some of the AEP mice have drastically higher cytokine levels than the age-matched controls, suggesting a progression of the disease. It has been reported that IFN-␥ is the key mediator in HLH development in mice (15), but we could not detect the same elevation in AEP-knockout mice. However, the report by Jordan et al. (15) that IFN-␥ is only essential for HLH development in a lymphocytic choriomeningitic virus (LCMV)induced fashion indicates that the IFN-␥ level might not necessarily be elevated in physiological condition in mice. In familial HLH, genetic studies mapped the HLH-associated genes in a variety of chromosome locations, including 9q21.3–22 (unknown gene), 10q21–22 (PRF1), 17q25 (UNC13D), 6q24 (STX11), 15q21(RAB27A), 1q42.1– 42.2 (LYST), and Xq25 (SH2D1A) (1). However, our screening of DNA sequence from 10 familial HLH patients revealed no mutation in the coding region of the AEP gene (data not shown). Depending on the racial and ethnic backgrounds of patients, the frequency of familial HLH caused by unknown genetic defect ranges from 17% to 70% (16). Therefore, more sequences from a larger pool of patients is necessary before we can exclude the possibility that a defect in AEP is implicated in HLH. Analysis of RBC membrane proteins on SDS/PAGE indicates that AEP plays an essential role for maintaining RBC membrane integrity. During the maturation of reticulocytes into erythrocytes, the membrane structures are remodeled such that a portion of the cell membrane is eliminated by fusion with a lysosome (17). Presumably, protease activities in AEP erythrocytes are impaired, leading to abnormal membrane protein destruction during the erythrocyte development. Indeed, activity of dipeptidyl peptidase II, a protease that can be found in erythrocytes in which activity depends on the maturation status (18), is dramatically elevated in Chan et al.

AEP-null tissues (19). The defective RBCs were actively engulfed by the histiocytes in the spleen and bone marrow, which might explain why the RBC count was evidently decreased in AEP-null mice. Nevertheless, EpoR was substantially elevated in the bone marrow from the AEP-null mice compared with wild-type mice. Concomitantly, Epo in blood was also markedly enhanced in AEP-deficient mice, which might be a reconciling mechanism for generating more mature RBCs in the anemic mice. Epo is primarily produced in the peritubular fibroblasts of the kidney in response to anemia, hypoxia, and, to a lesser extent, vasoconstriction (20). Epo may be nonphysiologically increased in association with kidney abnormalities (21). Previous study shows that AEP depletion incurs aberrant lysosomal storage in kidney-proximal tubule cells. Presumably, AEP-deficiency-provoked renal dysfunction provides another molecular mechanism for the overproduction of Epo in AEP-null mice. We observed that phagocytic activity of AEP-deficient macrophages against fluorescent microspheres and RBCs was slightly but significantly elevated, fitting the observation that numerous hemophagocytes were observed in AEP⫺/⫺ mice. However, the only slight enhancement of phagocytic activity of AEP-deficient macrophages does not account for the remarkable hemophagocytosis in AEP⫺/⫺ mice. One possibility is that phagocytic activity of Gaucher-like cells in the bone marrow and spleen, but not peritoneal macrophages, in AEP⫺/⫺ mice might be severely enhanced. Another possibility is that impaired digestive activity of macrophages against RBCs as well as enhanced phagocytic activity might cause the remarkable hemophagocytosis. As described, erythrocyte membrane integrity of AEP⫺/⫺ mice was impaired in an agedependent manner, suggesting that the defect in RBCs from AEP⫺/⫺ mice might cause hemophagocytosis in the bone marrow. However, circulating RBCs from AEP⫺/⫺ mice and ⫹/⫺ mice were equally engulfed by macrophages in the phagocytic assay. Engulfed RBCs in the bone marrow, but not circulating RBCs, might be morphologically impaired with membrane deformity. Conceivably, these morphologically aberrant RBCs might be engulfed in the bone marrow, so that they are not circulating in the peripheral tissues. Obviously, further characterization of Gaucherlike cells and RBCs in AEP⫺/⫺ mice is necessary to clarify the discrepancy. It has been proposed that the pathology of HLH largely depends on the failure of cytotoxic granule discharge in T cells. PNAS 兩 January 13, 2009 兩 vol. 106 兩 no. 2 兩 471

CELL BIOLOGY

Fig. 3. Extramedullary hematopoiesis of myeloid lineage in AEP-knockout mice. (A) Non-T and non-B cells increased in splenocytes with age and spleen weight in AEP-knockout mice. Spleen weight is shown above each graph. (B and C) A substantial increase of erythroid cells (Ter-119/CD45) (B) and small increase of myeloid cells (Gr-1/Mac-1) (C) occurred in spleen from AEP-null mice compared with wild-type mice.

A

E

B

C

D

Fig. 4. Enhanced phagocytic activity against RBCs in AEP-null macrophages. (A) H&E staining of the bone marrow and spleen from wild-type and AEP-lacking mice. AEP-null mice display ‘‘Gaucher-like cells’’ in bone marrow with engulfed RBCs. The bone marrow and spleen from AEP-null mice contained numerous Gaucher-like cells. The histiocytes in AEP-lacking mice contained numerous engulfed RBCs (arrows). (Scale bar, 10 ␮m.) (B) Electron microscopic analysis of histiocytes in bone marrow from AEP-null animals. R, RBC. (Scale bar, 2 ␮m.) (C) Enhanced phagocytic activity of peritoneal macrophages from AEP-knockout mice. Peritoneal macrophages from 2-month-old AEP-null mice (⫺/⫺) and control heterozygous mice (⫹/⫺) were used for the phagocytosis assay with fluorescent microspheres. (Left) Representative results. (Right) Results are shown as a histogram; data are expressed as mean ⫾ SD. ⴱ, P ⬍ 0.05, Student’s t test, n ⫽ 6. (D) Peritoneal macrophages from AEP-knockout mice (⫺/⫺) and control heterozygous mice (⫹/⫺) were used for the phagocytosis assay against RBCs from AEP-knockout mice and control heterozygous mice (⫹/⫺). RBCs were untreated (⫺) or opsonized (⫹) with anti-mouse RBC antibody before the assay. Macrophages (M␾) and RBCs were taken from 12-month-old mice and incubated for 1 h. Data are expressed as mean ⫾ SD. ⴱⴱⴱ, P ⬍ 0.001 against untreated group; ⴱⴱ, P ⬍ 0.001 against opsonized group in different genotype using the same RBCs, Student’s t test, n ⫽ 6. (E) Reduced natural killer cells activity in AEP-knockout mice. Percent cytotoxicity of NK cells against P815 mastocytoma from aged-matched wild-type (filled bar) and AEP-knockout (open bar) were measured by flow cytometry. Results are expressed as mean ⫾ SEM. ⴱ, P ⬍ 0.05, Student’s t test, n ⫽ 3.

This dependence prevents the ultimate execution of the antigenpresenting cells, and thus preventing the termination of the antigen presentation and T cell activation, leading to the excessive cytokine production and systemic inflammation (15). Although the role of AEP in cytotoxic processing remains unknown, it has been reported that AEP is essential for microbial tetanus toxin antigen presentation (9). Presumably, defective antigen presentation by AEP deficiency might be one of the causes for the hyperimmunological response in HLH. Materials and Methods Mice. AEP ⫹/⫹, ⫹/⫺, and ⫺/⫺ mice on a mixed 129/Ola and C57BL/6 background were generated as reported (19) and kept under specific pathogenfree conditions in an environmentally controlled clean room at the Whitehead Biomedical Research Building and Institute for Experimental Animals at Kanazawa University. Mice were housed at 22 °C on a 12-h/12-h light/dark cycle. Food and water were provided ad lib. The experiments were conducted according to the institutional ethical guidelines for animal experiments and approved by the Institutional Animal Care and Use Committee (IACUC) at Emory University and at Kanazawa University. Flow Cytometric Analysis. Single-cell suspensions of spleen were prepared by squeezing the spleen through a nylon mesh. Mature erythrocytes were removed by lysis in pH 7.4 Tris-buffered hypotonic ammonium chloride solution (144 mM). Directly conjugated monoclonal antibodies to Ter119, CD45, Gr-1, Mac-1, CD3, and B220 (all from BD Biosciences PharMingen) were used to stain the cells. Nonviable cells were excluded from analysis by staining with 7-aminoactinomycin D (BD Biosciences PharMingen) or propidium iodide (Sigma). A 472 兩 www.pnas.org兾cgi兾doi兾10.1073兾pnas.0809824105

four-color FACSCalibur cytometer (BD Biosciences) was used to acquire a minimum of 10,000 events. Data were analyzed with FloJo software (Treestar). Serum Epo Determination. Mouse serum Epo levels were determined by using the Erythropoietin Quantikine ELISA Kit (R&D Systems). Immunohistochemistry Staining and Histological Examination. Immunohistochemical assays were performed on formalin-fixed paraffin-embedded sections. Three- to 5-␮m thick sections from femur bone marrow and the spleen were cut, deparaffinized in xylene, rehydrated in graded alcohols, and subjected to hematoxylin and eosin (H&E) or periodic acid–Schiff (PAS) staining using standard methods. The autofluorescence of bone marrow cells was analyzed by confocal microscopy (LCM5, Carl Zeiss). For cytospins, spleen and bone marrow cell suspensions were prepared, spun onto slides, and stained with Giemsa stain. For immunohistochemical analysis, the slides were treated with 0.3% hydrogen peroxide and SuperBlock blocking buffer (Pierce) and incubated with mAb against F4/80 (Abcam) and CD68 (Serotec). Next, they were treated with biotinconjugated Ab against rat IgG (Vector Laboratories), and then with an ABC kit (Vector Laboratories). Finally, horseradish peroxidase activity was detected by DAB substrate and the cells were counterstained with hematoxylin. For Epo receptor staining, endogenous peroxidase activity was blocked by 3% hydrogen peroxide for 5 min and all slides were boiled in 10 mM sodium citrate buffer (pH 6.0) for 10 min. EpoRs on the tissues were stained by using a Zymed Histo-SP AEC kit (Invitrogen) and specific antibody against EpoR (sc-697, 1:100 dilution, Santa Cruz Biotechnology). Slides were then counterstained with hematoxylin. The slides were examined under an Olympus BX41 microscope. RBC Membrane Protein Analysis. RBC membrane was isolated as previously described (22). Purified RBC ghosts were then solubilized, resolved by SDS/

Chan et al.

Electron Microscopy. Mice were fixed by injecting 2.5% glutaraldehyde into the heart, and the femurs were decalcified by 10% EDTA; the bone marrows were subjected to standard electron microscopic techniques as described previously (19, 23). Ultrathin sections were observed under a Hitachi H7600 electron microscope. Phagocytosis Assay. Peritoneal macrophages were harvested on day 4 after intraperitoneal injection of 1.5 ml of 3% thioglycollate (Difco). Macrophages (2 ⫻ 105 cells) were plated on round 12-mm glass coverslips on a 24-well plate. Macrophages were incubated with FluoSpheres fluorescent microsphere (1 ␮m, 2 ⫻ 107 beads, Molecular Probes) for 2 h, or RBCs (4 ⫻ 106 cells) for 1 h or 24 h at 37 °C. After incubation, the cells were washed out to remove surfacebound beads. For erythrophagocytosis assay, unengulfed RBCs were removed by hypotonic lysis, fixed, and stained with Giemsa stain. Engulfed microspheres and RBCs were observed by fluorescence and light microscopy, respectively. Phagocytic index is expressed as the number of engulfed microspheres or RBCs per macrophage.

were centrifuged and the RBCs were lysed in ACK buffer (150 mM NH4Cl, 1 mM KHCO3, and 0.1 mM EDTA, pH 7.2) at room temperature. The spleen suspension was then incubated with P815 murine mastocytoma prelabeled with Vybrant cell labeling solution (Invitrogen) at an effector/target ratio of 50:1. The mixture was incubated at 37 °C for 3 h. Fifteen microliters of propidium iodide (50 ␮g/ml) was added to the effector/target cell mixture during incubation. Natural killer cell functions were analyzed by flow cytometry as reported (24). Statistics. Data are presented as mean ⫾ SEM or S.D. as indicated in the figure legends. Statistical evaluation was carried out by means of Student’s t test or Welch’s t test following Levene’s test for equality of variance between AEPdeficient and control mice. A two-sided level of P ⬍ 0.05 was accepted as statistically significant. All statistical analysis was performed by the computer program Prism (GraphPad).

NK Cell Activity Assay. Spleen cell suspensions were prepared by squeezing the organs through a nylon mesh (100 ␮m) in DMEM containing 5% FBS. The cells

ACKNOWLEDGMENTS. We thank the late Dr. O. Miyaishi of Chubu Rosai Hospital for his initial contribution on this work and Drs. A. Shiratsuchi and Y. Nakanishi at Kanazawa University for teaching us the phagocytic assay. This work is supported by Grant R01 NS45627 from the National Institutes of Health (to K.Y.) and Grant 18500325 from the Ministry of Education, Culture, Sports, Science and Technology of Japan (to N.H.).

1. Janka GE (2007) Familial and acquired hemophagocytic lymphohistiocytosis. Eur J Pediatr 166:95–109. 2. Janka GE (2007) Hemophagocytic syndromes. Blood Rev 21:245–253. 3. Stepp SE, et al. (1999) Perforin gene defects in familial hemophagocytic lymphohistiocytosis. Science 286:1957–1959. 4. Feldmann J, et al. (2003) Munc13-4 is essential for cytolytic granules fusion and is mutated in a form of familial hemophagocytic lymphohistiocytosis (FHL3). Cell 115:461– 473. 5. Menasche G, et al. (2000) Mutations in RAB27A cause Griscelli syndrome associated with haemophagocytic syndrome. Nat Genet 25:173–176. 6. Nagle DL, et al. (1996) Identification and mutation analysis of the complete gene for Chediak-Higashi syndrome. Nat Genet 14:307–311. 7. Chen JM, Dando PM, Stevens RA, Fortunato M, Barrett AJ (1998) Cloning and expression of mouse legumain, a lysosomal endopeptidase. Biochem J 335:111–117. 8. Chen JM, Fortunato M, Barrett AJ (2000) Activation of human prolegumain by cleavage at a C-terminal asparagine residue. Biochem J 352:327–334. 9. Manoury B, et al. (1998) An asparaginyl endopeptidase processes a microbial antigen for class II MHC presentation. Nature 396:695– 699. 10. Manoury B, et al. (2002) Destructive processing by asparagine endopeptidase limits presentation of a dominant T cell epitope in MBP. Nat Immunol 3:169 –174. 11. Liu Z, et al. (2008) Neuroprotective actions of PIKE-L by inhibition of SET proteolytic degradation by asparagine endopeptidase. Mol Cell 29:665– 678. 12. Sasaki R, Masuda S, Nagao M (2000) Erythropoietin: Multiple physiological functions and regulation of biosynthesis. Biosci Biotechnol Biochem 64:1775–1793. 13. Henter JI, Arico M, Elinder G, Imashuku S, Janka G (1998) Familial hemophagocytic lymphohistiocytosis. Primary hemophagocytic lymphohistiocytosis. Hematol Oncol Clin North Am 12:417– 433.

14. Janka G, Imashuku S, Elinder G, Schneider M, Henter JI (1998) Infection- and malignancy-associated hemophagocytic syndromes. Secondary hemophagocytic lymphohistiocytosis. Hematol Oncol Clin North Am 12:435– 444. 15. Jordan MB, Hildeman D, Kappler J, Marrack P (2004) An animal model of hemophagocytic lymphohistiocytosis (HLH): CD8⫹ T cells and interferon gamma are essential for the disorder. Blood 104:735–743. 16. Zur Stadt U, et al. (2006) Mutation spectrum in children with primary hemophagocytic lymphohistiocytosis: Molecular and functional analyses of PRF1, UNC13D, STX11, and RAB27A. Hum Mutat 27:62– 68. 17. Gronowicz G, Swift H, Steck TL (1984) Maturation of the reticulocyte in vitro. J Cell Sci 71:177–197. 18. Maes MB, Scharpe S, De Meester I (2007) Dipeptidyl peptidase II (DPPII), a review. Clin Chim Acta 380:31– 49. 19. Shirahama-Noda K, et al. (2003) Biosynthetic processing of cathepsins and lysosomal degradation are abolished in asparaginyl endopeptidase-deficient mice. J Biol Chem 278:33194 –33199. 20. Fisher RC, et al. (2004) PU.1 supports proliferation of immature erythroid progenitors. Leuk Res 28:83– 89. 21. Grotta JC, Manner C, Pettigrew LC, Yatsu FM (1986) Red blood cell disorders and stroke. Stroke 17:811– 817. 22. Marchesi VT, Palade GE (1967) The localization of Mg-Na-K-activated adenosine triphosphatase on red cell ghost membranes. J Cell Biol 35:385– 404. 23. Koh S, et al. (1993) Immunoelectron microscopic localization of the HPC-1 antigen in rat cerebellum. J Neurocytol 22:995–1005. 24. Kane KL, Ashton FA, Schmitz JL, Folds JD (1996) Determination of natural killer cell function by flow cytometry. Clin Diagn Lab Immunol 3:295–300.

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10% PAGE, and subjected to silver staining. Protein bands of interest were excised from the gel and analyzed by proteomic analysis.

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BIOCHEMISTRY

Correction for ‘‘Mice lacking asparaginyl endopeptidase develop disorders resembling hemophagocytic syndrome,’’ by Chi-Bun Chan, Michiyo Abe, Noriyoshi Hashimoto, Chunhai Hao, Ifor R. Williams, Xia Liu, Shinji Nakao, Akitsugu Yamamoto, Shi-Yong Li, Ikuko Hara-Nishimura, Masahide Asano, and Keqiang Ye, which appeared in issue 2, January 13, 2009, of Proc Natl Acad Sci USA (106:468–473; first published December 23, 2008; 10.1073兾pnas.0809824105). The authors request that Chengyun Zheng, Jan-Inge Henter, Marie Meeths, and Magnus Nordenskjold be added to the authors list between Akitsugu Yamamoto and Shi-Yong Li. Chengyun Zheng designed research and analyzed data, Jan-Inge Henter designed research, Marie Meeths performed research and analyzed data, and Magnus Nordenskjold analyzed data. The online version has been corrected. The corrected author and affiliation lines and related footnotes appear below.

Correction for ‘‘Crystal structure of a near-full-length archaeal MCM: Functional insights for an AAA⫹ hexameric helicase,’’ by Aaron S. Brewster, Ganggang Wang, Xian Yu, William B. Greenleaf, Jose´ Marı´a Carazo, Matthew Tjajadia, Michael G. Klein, and Xiaojiang S. Chen, which appeared in issue 51, December 23, 2008, of Proc Natl Acad Sci USA (105:20191– 20196; first published December 10, 2008; 10.1073/ pnas.0808037105). The authors note that the author name Matthew Tjajadia should have appeared as Matthew Tjajadi. The online version has been corrected. The corrected author line appears below.

aDepartment of Pathology and Laboratory Medicine, Emory University School of Medicine, 615 Michael Street, Atlanta, GA 30322; bDivision of Transgenic Animal Science, Advanced Science Research Center, and cDepartment of Cellular Transplantation Biology, Graduate School of Medical Science, Kanazawa University, 13-1 Takara-machi, Kanazawa 920-8640, Japan; dDepartment of Bio-Science, Nagahama Institute of Bio-Science and Technology, 1266 Tamura-machi, Nagahama 526-0829, Japan; gDepartment of Botany, Graduate School of Science, Kyoto University, Kitashirakawa, Sakyo-ku, Kyoto 606-8502, Japan; eDepartment of Woman and Child Health, Childhood Cancer Research Unit, and fDepartment of Molecular Medicine and Surgery, Clinical Genetics Unit, Karolinska Institute, Karolinska University Hospital, SE-141 86 Stockholm, Sweden

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NEUROSCIENCE

Correction for ‘‘Caloric restriction improves memory in elderly humans,’’ by A. V. Witte, M. Fobker, R. Gellner, S. Knecht, and A. Flo ¨el, which appeared in issue 4, January 27, 2009, of Proc Natl Acad Sci USA (106:1255–1260; first published January 26, 2009; 10.1073/pnas.0808587106). The authors note that the following two sentences should have been included on page 1257, left column, first full paragraph, after the sentence ending in the fifth line: ‘‘Note that one subject had to be excluded from the correlation analysis for insulin and one subject had to be excluded for hs-CRP due to a measurement error in insulin or hs-CRP, respectively. Therefore, analyses were conducted with 9 of initially 10 subjects defined by weight loss of ⬎2 kg.’’ This error does not affect the conclusions of the article. www.pnas.org兾cgi兾doi兾10.1073兾pnas.0901549106

Author contributions: C.-B.C., M. Abe, N.H., C.H., I.R.W., C.Z., J.-I.H., M. Asano, and K.Y. designed research; C.-B.C., M. Abe, N.H., C.H., I.R.W., X.L., S.N., A.Y., and M.M. performed research; I.H.-N. contributed new reagents/analytic tools; C.-B.C., M. Abe, N.H., C.H., I.R.W., C.Z., M.M., M.N., S.-Y.L., M. Asano, and K.Y. analyzed data; and C.-B.C., M. Asano, and K.Y. wrote the paper. 1C.-B.C.

and M. Abe contributed equally to this work.

2To

whom correspondence may be addressed. E-mail: [email protected] or [email protected].

www.pnas.org兾cgi兾doi兾10.1073兾pnas.0901254106

PNAS 兩 April 7, 2009 兩 vol. 106 兩 no. 14 兩 6023

CORRECTIONS

Chi-Bun Chana,1, Michiyo Abeb,1, Noriyoshi Hashimotob, Chunhai Haoa, Ifor R. Williamsa, Xia Liua, Shinji Nakaoc, Akitsugu Yamamotod, Chengyun Zhenge,f, Jan-Inge Hentere, Marie Meethse,f, Magnus Nordenskjoldf, Shi-Yong Lia, Ikuko Hara-Nishimurag, Masahide Asanob,2, and Keqiang Yea,2

Aaron S. Brewster, Ganggang Wang, Xian Yu, William B. Greenleaf, Jose´ Marı´a Carazo, Matthew Tjajadi, Michael G. Klein, and Xiaojiang S. Chen

Correction

CELL BIOLOGY

Correction for ‘‘Mice lacking asparaginyl endopeptidase develop disorders resembling hemophagocytic syndrome,’’ by Chi-Bun Chan, Michiyo Abe, Noriyoshi Hashimoto, Chunhai Hao, Ifor R. Williams, Xia Liu, Shinji Nakao, Akitsugu Yamamoto, Shi-Yong Li, Ikuko Hara-Nishimura, Masahide Asano, and Keqiang Ye, which appeared in issue 2, January 13, 2009, of Proc Natl Acad Sci USA (106:468–473; first published December 23, 2008; 10.1073兾pnas.0809824105). The authors request that Chengyun Zheng, Jan-Inge Henter, Marie Meeths, and Magnus Nordenskjold be added to the authors list between Akitsugu Yamamoto and Shi-Yong Li. Chengyun Zheng designed research and analyzed data, Jan-Inge Henter designed research, Marie Meeths performed research and analyzed data, and Magnus Nordenskjold analyzed data. The online version has been corrected. The corrected author and affiliation lines and related footnotes appear below.

CORRECTION

Chi-Bun Chana,1, Michiyo Abeb,1, Noriyoshi Hashimotob, Chunhai Haoa, Ifor R. Williamsa, Xia Liua, Shinji Nakaoc, Akitsugu Yamamotod, Chengyun Zhenge,f, Jan-Inge Hentere, Marie Meethse,f, Magnus Nordenskjoldf, Shi-Yong Lia, Ikuko Hara-Nishimurag, Masahide Asanob,2, and Keqiang Yea,2 aDepartment of Pathology and Laboratory Medicine, Emory University School of Medicine, 615 Michael Street, Atlanta, GA 30322; bDivision of Transgenic Animal Science, Advanced Science Research Center, and cDepartment of Cellular Transplantation Biology, Graduate School of Medical Science, Kanazawa University, 13-1 Takara-machi, Kanazawa 920-8640, Japan; dDepartment of Bio-Science, Nagahama Institute of Bio-Science and Technology, 1266 Tamura-machi, Nagahama 526-0829, Japan; gDepartment of Botany, Graduate School of Science, Kyoto University, Kitashirakawa, Sakyo-ku, Kyoto 606-8502, Japan; eDepartment of Woman and Child Health, Childhood Cancer Research Unit, and fDepartment of Molecular Medicine and Surgery, Clinical Genetics Unit, Karolinska Institute, Karolinska University Hospital, SE-141 86 Stockholm, Sweden

Author contributions: C.-B.C., M. Abe, N.H., C.H., I.R.W., C.Z., J.-I.H., M. Asano, and K.Y. designed research; C.-B.C., M. Abe, N.H., C.H., I.R.W., X.L., S.N., A.Y., and M.M. performed research; I.H.-N. contributed new reagents/analytic tools; C.-B.C., M. Abe, N.H., C.H., I.R.W., C.Z., M.M., M.N., S.-Y.L., M. Asano, and K.Y. analyzed data; and C.-B.C., M. Asano, and K.Y. wrote the paper. 1C.-B.C.

and M. Abe contributed equally to this work.

2To

whom correspondence may be addressed. E-mail: [email protected] or [email protected].

www.pnas.org兾cgi兾doi兾10.1073兾pnas.0901254106

www.pnas.org

PNAS 兩 April 7, 2009 兩 vol. 106 兩 no. 14 兩 6023