extfieldsofstudy: Medicine Biology

text:
In Situ Immunolocalization and Stage-Dependent Expression of a Secretory Serine Protease in Leishmania donovani and Its Role as a Vaccine Candidate

ABSTRACT Proteases have been found to play essential roles in many biological processes, including the pathogenesis of leishmaniasis. Most parasites rely on their intracellular and extracellular protease repertoire to invade and multiply in mammalian host cells. However, few studies have addressed serine proteases in Leishmania and their role in host pathogenesis. Here we report the intracellular distribution of a novel L. donovani secretory serine protease in the flagellar pocket, as determined by immunogold labeling. Flow cytometry and confocal immunofluorescence analysis revealed that the expression of the protease diminishes sequentially from virulent to attenuated strains of this species and is also highly associated with the metacyclic stage of L. donovani promastigotes. The level of internalization of parasites treated with the anti-115-kDa antibody into host macrophages was significantly reduced from that of non-antibody-treated parasites, suggesting that this serine protease probably plays a role in the infection process. In vivo studies confirmed that this serine protease is a potential vaccine candidate. Altogether, the 115-kDa serine protease might play vital roles in L. donovani pathogenesis and hence could be recognized as a potential candidate for drug design.

Leishmania species, belonging to the family Trypanosomatidae, are a group of protozoan pathogens that cause a spectrum of chronic diseases ranging from self-healing cutaneous lesions to a lethal visceral disorder. They represent a major health problem in tropical and subtropical areas around the world. Leishmania species have a relatively simple life cycle, with parasite stage differentiation regulated by environmental signals encountered in their different hosts. They are digenetic parasites: the flagellated promastigote forms, which can be derived in axenic culture, differentiate within the alimentary tracts of sandfly vectors from a replicating procyclic to a nonreplicating, infectious metacyclic stage, whereas obligately intracellular amastigotes live and replicate in mammalian mononuclear phagocytes, such as macrophages (1).
Recent advances in genomic analysis of several of the major global parasites have revealed key factors involved in the pathogenesis of parasite diseases. Among the major virulence factors identified are parasite-derived proteases. Parasite proteases play significant roles in the pathogenesis of parasitic diseases; the proteases are involved in the invasion of the host via parasite migration through tissue barriers, degradation of host proteins for their nutrition, immune evasion, and activation of inflammation (21). The migration of the parasite in host tissues is mediated by the release of proteolytic enzymes that can degrade the tissue barriers. Potent proteolytic enzymes of cysteine, serine, and metalloprotease classes have been identified in secretory products of many of the parasites (8,19,26,27,30). Serine proteases have been reported to have strong hydrolytic activity against a wide range of extracellular matrix (ECM) components and human plasma proteins; hence, they are thought to play vital roles in the host tissue invasion process (17). These proteases are found to localize in different cellular compartments. In trypanosomatids, the flagellar pocket, a specialized region formed by invagination of the plasma membrane, has been found to harbor a virulence-associated protease destined to be exported extracellularly (11,31). However, the specific molecular function of the extracellular serine protease of the Indian strain of Leishmania donovani has not been elucidated yet.
Our previous study demonstrated that a 115-kDa serine protease is secreted by an Indian strain of L. donovani (8). In the present report, we have demonstrated the intracellular localization of this parasite-derived secreted serine protease and also its differential expression in virulent, avirulent, and attenuated strains of L. donovani as well as in different cell cycle stages of the parasite. A combination of in vitro and in vivo experiments suggests that the protease might play a role in host-parasite interactions and could be validated as a potential vaccine candidate.

MATERIALS AND METHODS
Culture of L. donovani and macrophages. Leishmania donovani strain MHOM/ IN/1983/AG83 was isolated from Indian patients with visceral leishmaniasis. The promastigotes used were at or near the stationary phase of growth in the 4th passage (4th P) (virulent) or 34th passage (34th P) (avirulent) of in vitro culture after transformation from liver-or spleen-derived amastigotes. Promastigotes were cultured at 22°C in medium 199 with Hanks' salt containing HEPES, L-glutamine, 10% heat-inactivated fetal calf serum (FCS), penicillin at 50 U ml Ϫ1 , and streptomycin at 50 g ml Ϫ1 (all from Gibco BRL/Life Technologies, Middlesex, United Kingdom) (24). Amastigotes of Leishmania donovani were isolated from the spleens of infected mice by homogenization of the organs, followed by Percoll gradient fractionation, as described elsewhere (6). The axenic amastigotes were cultured according to the method of Li et al. (20). Axenic amastigote cells were visualized under a light microscope to confirm transfor-mation; these were harvested 4 days after initial differentiation, with at least one serial dilution. UR6 promastigotes of L. donovani were cultured on solid blood agar slants containing glucose, sodium chloride, peptone, beef extract, and rabbit blood supplemented with gentamicin (7). UR6 cells were harvested by scraping in 10 mM phosphate-buffered saline (PBS) (pH 7.2) from 3-day-old blood agar slants. For subsequent flow cytometry experiments, UR6 cells were maintained in liquid medium (medium 199) supplemented with 10% FCS. Metacyclic parasites of virulent and attenuated cells were isolated from 2-and 4-day-old cultures by the peanut agglutinin (PNA) agglutination method (29).
The murine macrophage cell line RAW 264.7 was cultured at 37°C under 5% CO 2 in RPMI 1640 supplemented with 10% heat-inactivated FCS, 10 mM HEPES (pH 7.3), 100 U/ml penicillin, and 100 g/ml streptomycin. For experiments, cells between passages 3 and 10 were seeded on sterile glass coverslips and were grown overnight.
Identification and purification of the 115-kDa serine protease. An extracellular serine protease with a molecular mass of 115 kDa from an Indian strain of Leishmania donovani has been reported (8). Briefly, the extracellular proteases secreted in the culture supernatant of early-passage (4th-passage) L. donovani were subjected to gelatin substrate gel electrophoresis and were treated with different classes of protease inhibitors. We noted that the activity of the 115-kDa protease was completely inhibited by the serine protease inhibitor aprotinin. The protease was then purified by ammonium sulfate precipitation and was subjected to aprotinin-agarose affinity chromatography, followed by continuous elution electrophoresis. The purified serine protease was further characterized (8).
Raising of antisera. A polyclonal antibody against the 115-kDa extracellular protease was raised by subcutaneous injection of 80 g of purified protease emulsified in Freund's complete adjuvant into a male rabbit. Three booster doses were administered at intervals of 2 weeks by injecting the purified protease emulsified in Freund's incomplete adjuvant. At 10 days after the fourth injection, blood was collected from the rabbit's ear, and the antiserum against the 115-kDa protease was separated (25). A mouse polyclonal antiserum was also raised against purified L. donovani gp63, which was purified according to the method described by Soteriadou et al. (32). A polyclonal antibody was purified from the antiserum by protein A agarose (Sigma Chemical Co., St. Louis, MO) affinity chromatography (14). The peak fractions were concentrated and kept frozen at Ϫ80°C.
Electrophoresis and immunoblotting. Western blotting was carried out according to the method described by Towbin et al. (33) with some modifications. Briefly, cell-free cultured supernatants of virulent promastigotes and whole-cell lysates of virulent promastigotes, avirulent promastigotes, and attenuated UR6 cells were resolved by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) (18) under nonreducing conditions and were transferred to nitrocellulose membranes (pore size, 0.45 m). Membranes were incubated for 1 h with the anti-115-kDa polyclonal antiserum (dilution, 1: 500) in Tris-buffered saline (TBS) at 22°C, washed in TBS, and incubated with appropriate horseradish peroxidase (HRP)-conjugated secondary antibodies. Membranes were developed with 0.2 mM 4-chloro-1-naphthol.
Flow cytometric analysis. The anti-115-kDa extracellular protease antibody was labeled with fluorescein isothiocyanate (FITC) by using the Pierce EZ-Label FITC protein labeling kit. Early-passage (4th-P), late-passage (34th-P), and UR6 cells and axenic amastigotes of virulent AG83 were harvested and stained with the FITC-labeled anti-115-kDa extracellular protease antibody. Procyclic and metacyclic promastigotes of early-passage virulent cells and attenuated cells from 3-and 4-day-old cultures were also isolated and stained with the FITC-labeled anti-115-kDa extracellular protease antibody. Briefly, the cells were washed three times in PBS and were then fixed in 1% paraformaldehyde. The fixed cells were taken up in a fluorescence-activated cell sorter (FACS) buffer (catalog no. 333138; Becton Dickinson) and were stained with the FITC-labeled anti-115-kDa protease (dilution, 1:100). The fluorescence of the fixed stained cells was detected on a FACSCalibur system (BD Biosciences), and the data were analyzed using CellQuest software.
Confocal immunofluorescence microscopy. L. donovani promastigotes (early and late passages) and UR6 cells were centrifuged at 3,000 ϫ g for 10 min, washed in PBS, and resuspended at a concentration of 10 7 cells/ml. Cells were applied to poly-L-lysine-coated glass coverslips and were fixed in PBS containing 4% paraformaldehyde for 30 min. Coverslips were rinsed once with PBS to remove the fixative and were incubated in a blocking solution consisting of PBS supplemented with 5% FCS, 0.1% Tween 20, and 0.1% Triton X-100 for 1 h at room temperature. To investigate the specificity of the anti-115-kDa antibody, an irrelevant antibody (anti-gp63) was tested. A primary antibody against L. donovani gp63 (raised in mouse) and a purified antibody against the L. donovani 115-kDa serine protease (raised in rabbit) were diluted 1:500 in blocking solution and were incubated separately or together with the promastigotes of L. donovani.
After incubation of fixed parasites with the primary antibody for 1 h, coverslips were rinsed six times for 5 min with a PBS wash solution containing 0.1% Tween 20. Fixed cells were then incubated separately or together with an FITC-conjugated anti-rabbit secondary antibody at a 1:1,000 dilution and with a tetramethyl rhodamine isocyanate (TRITC)-conjugated anti-mouse secondary antibody at a 1:100 dilution in blocking buffer. The incubation and all subsequent steps were performed in the dark. Secondary antibodies were removed by soaking coverslips six times, for 5 min each time, in wash buffer. Cells were then washed in PBS and were observed under a laser scanning microscope (LSM 510; Carl Zeiss, Jena, Germany). The images were further processed using Adobe Photoshop, version 7.0 (Adobe Systems).
Immunogold labeling. Approximately 10 7 early-passage promastigotes and amastigotes of L. donovani were fixed in 2% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.4) for 4 h at 4°C. Samples were dehydrated in ethanol and embedded in LR White resin. Ultrathin sections were cut and incubated with an anti-115-kDa extracellular serine protease antibody (1:20). The binding of the primary antibody was visualized by incubation with a gold (diameter, 10 nM)conjugated secondary antibody (1:50). Samples were stained with uranyl acetate and with lead citrate and were finally investigated with an FEI Tecnai G 2 transmission electron microscope (with the SIS Protem program) (15).
Effects of antisera on macrophage infectivity. Parasites were incubated for 1 h with the anti-115-kDa antibody (dilution, 1:200) and were then washed three times with excess PBS to remove the unbound antibody completely. In parallel experiments, parasites were also treated with preimmune serum. Preimmune serum-or antibody-treated promastigotes and untreated promastigotes were used to infect cultures of adherent macrophages (RAW 264.7 cell line) on glass coverslips (1 ϫ 10 6 macrophages/coverslip) in 0.5 ml of RPMI 1640 contain-ing10% FCS at a ratio of 10 parasites per macrophage. The coverslips were kept at 37°C under a humidified 5% CO 2 atmosphere. After sequential incubation for 24 h, 48 h, and 72 h, the cells were fixed in methanol and were stained with Giemsa stain for the determination of intracellular parasite numbers (2,22). In a separate set of experiments, parasites pretreated with the anti-115-kDa antibody were incubated with the macrophages, which had previously been incubated only with the purified 115-kDa protease (1 g/ml) (4), and the experiments proceeded as described above.
In vivo assay of L. donovani infectivity after immunization with the protease. Groups of six female BALB/c mice, 4 to 6 weeks old, were vaccinated subcutaneously or intraperitoneally with 10 g of the purified 115-kDa serine protease in Freund's complete adjuvant in a volume of 0.1 ml. Three booster doses were given in Friend's incomplete adjuvant at biweekly intervals with the same amount of protein. Control mice received the adjuvant alone. One month after the last booster, one set of mice was challenged intracardially with 5 ϫ 10 5 virulent L. donovani promastigotes, and another set of mice was challenged with 5 ϫ 10 8 parasites. Starting at 4 weeks postchallenge, the mice were sacrificed at different times up to 8 months, and parasite burdens in the spleen and liver were quantitatively evaluated (5) by microscopic observation of Giemsa-stained preparations of impression smears of the tissues.

RESULTS AND DISCUSSION
Immunoblotting. A polyclonal antibody raised against the purified serine protease was employed in Western blot analysis of the whole-cell lysates of 4th-P, 34th-P, and UR6 promastigotes. The cell-free culture supernatant of the early-passage cell was analyzed by immunoblotting as well. It was observed in the experiment that a 115-kDa band appeared in all the lanes, but the intensities of the bands differed significantly (Fig. 1). The prominent band observed in Fig. 1, lane b, unambiguously suggested the intracellular appearance of the protease in virulent L. donovani promastigotes. Moreover, Western blotting for detection of the protease in whole-cell extracts of 34th-P L. donovani promastigotes (Fig. 1, lane c)  In the present study, flow cytometric analysis suggested a negative correlation between the level of expression of the 115-kDa protein and the number of times the parasites were cultivated in vitro, and hence a positive correlation with their virulence. As observed in Fig. 2I, the anti-protease antibody recognized 38.49% of the cells of the virulent 4th-P promastigotes and only 20.52% of the avirulent 34th-P promastigotes, suggesting reduced expression of the protein with an increase in serial cell passage number during in vitro cultivation, when the parasites also exhibit a relative loss of virulence. The anti-115-kDa antibody was found to recognize only 18.24% of the nonpathogenic attenuated UR6 promastigotes (Fig. 2I), as determined with reference to the number of control cells recognized, whereas the same antibody reacted with 18.36% of the total population of infective amastigotes. This lesser recognition of amastigotes, as well as of avirulent and attenuated promastigotes, might be due to the lower expression of the protease in these cells. These observations indicate that the 115-kDa serine protease is an exclusive secretory product of virulent L. donovani promastigotes, rather than of avirulent and attenuated promastigotes or infective amastigotes of virulent L. donovani. These results also demonstrate that a discrete population of cells is associated with abundant expression of 115-kDa protease. The growth of Leishmania promastigotes within the sandfly midgut or in in vitro culture is accompanied by differentiation of some of these organisms into metacyclic-stage promastigotes, which can be distinguished from noninfective procyclic promastigotes by their loss of agglutination by PNA lectin (29). The metacyclic stages of promastigotes are uniquely adapted to survival within vertebrate hosts. On the other hand, the relative loss of virulence of PNA-agglutinable (PNA ϩ ) procyclic promastigotes could suggest considerable differences in the metacyclogenetic potential of the parasites with different lengths of time in culture. Thus, both noninfective and infective stages of Leishmania promastigotes within the cultured populations are associated with certain specific developmental changes that could help this parasite to adapt to, and survive within, the vertebrate host. Differentiation of procyclic promastigotes into metacyclic promastigotes (metacyclogenesis) is known to be crucial for infectivity (3,9). Infective-stage promastigotes are generated in vitro as organisms approach the stationary phase of growth (2,29). To correlate these changes in the expression of the 115-kDa protease with metacyclogenesis, the expression of the protease was studied in promastigotes derived from 0 to 96 h of culture. Increased infectivity of virulent L. donovani promastigotes from early-log phase to stationary phase was observed with the increase in the percentage of PNA Ϫ promastigotes from 0 to 96 h (data not shown), and concurrently, the expression of the 115-kDa protease was found to be high, reaching a plateau at 96 h (Fig. 2II). da Silva and Sacks (9) have also shown that log-phase promastigotes represent a homogeneous population of noninfective parasites, whereas as many as 50% of the stationary-phase organisms appeared to be transformed into infective-stage promastigotes. To further confirm the association of the protease with virulence, the expression of the protease was studied in metacyclic (PNA Ϫ ) and procyclic (PNA ϩ ) promastigotes isolated from 12-, 24-, 48-, 72-, and 96-h cultures. It was noted that the expression of the protease was strongly associated with metacyclic promastigotes. Representative results for the expression of the 115-kDa protease in metacyclic and procyclic promastigotes isolated from 72-h and 96-h cultures of virulent and attenuated strain are shown in Fig. 2III. The results of this study showed 38.23% (72 h) and 38.97% (96 h) recognition in metacyclic promastigotes of 4th-passage L. donovani by the anti-115-kDa protease antibody, compared to 18.21% (72 h) and 19.01% (96 h) in procyclic promastigotes of the same early-passage promastigotes. For attenuated UR6 promastigotes, all of which were found to be PNA ϩ , the anti-115-kDa protease antibody was found to recognize 17.27% (72 h) and 18.16% (96 h) of cells of the total population (Fig. 2III).
During metacyclogenesis, several proteins with functions in protein synthesis, protein metabolism, and protein folding, etc., are downregulated because these promastigotes are in stationary phase and are mostly growth arrested (23). But Gull (13) found an increased abundance of isoforms of proteins involved in motility, including paraflagellar rod protein, ␣-tubulin, and ␤-tubulin, in metacyclic promastigotes, which are more active and motile. Similarly, in the present study, the increased expression and secretion of the 115-kDa protease in the infective metacyclic promastigotes could suggest an active role of this protease in the invasion process. Thus, the presence of the 115-kDa serine protease in the culture medium of virulent promastigotes of L. donovani could be a significant phenotypic marker of relative virulence displayed by the promastigotes obtained from culture.
Confocal microscopy. We used confocal immunofluorescence microscopy to compare the relative intensities and hence the relative expression levels of the 115-kDa protease in the virulent and the nonpathogenic attenuated promastigotes of an Indian strain of L. donovani. Cells were first incubated in the presence of a primary antibody against the serine protease; thereafter, they were probed with a fluorescence-conjugated secondary antibody, and intensities were visualized by confocal immunofluorescence microscopy. In order to confirm the localization and the specificity of recognition of the 115-kDa anti-protease antibody, confocal microscopy was also performed with an antibody against Leishmania gp63. In the immunofluorescence assay, the anti-115-kDa antibody was found to react in the cell body, where labeling was restricted to the posterior region of the promastigotes. By comparison of the relative intensities of the immunolabeled protease (Fig. 3), it appears that the relative quantity of the 115-kDa protein is directly proportional to the virulence of the parasite. This is in agreement with the fact that early-passage, virulent promastigotes (Fig. 3A2) showed significantly more-intense labeling than late-passage, avirulent promastigotes (Fig. 3B2), a finding consistent with the in vitro flow cytometric expression pattern. Similarly, the intensity of expression of the protease in the nonpathogenic attenuated UR6 cells (Fig. 3C2) was extremely low. Figure 3A3, B3, and C3 (red channel) are immunoconfocal images of parasites treated with the anti-gp63 antibody.
The images of the green and red channels were merged into single images (Fig. 3A4, B4, and C4), where yellow fluorescence represents the colocalization of the red and green fluorescence signals. The phase-contrast images of the same fields are shown in Fig. 3A1, B1, and C1. No fluorescence was detected in parallel preparations probed with the preimmune serum (Fig. 3D, E, and F). These results corroborate our findings that the increased level of the 115-kDa serine protease in early-passage, virulent parasites over that in late-passage, avirulent parasites was quite significant. Hence, this direct re- lationship of parasite virulence to protease expression suggests that the protease might play some role in host infection. Work is in progress in our laboratory to determine the exact role of this protease in pathogenesis. Immunoelectron microscopy. To gain insight into the precise cellular distribution of the serine protease, we employed an immunogold electron microscopy technique. Electron micrographs of thin sections of amastigotes and early-passage promastigotes revealed that most of the labeling seems to be associated with the flagellar pocket region at both stages of the parasite (Fig. 4). This finding clearly indicates that the 115-kDa protease is distributed throughout the flagellar pocket region, a compartment in protozoan architecture that is formed by an invagination of the plasma membrane where the flagellum emerges from the cell body. In recent years, it has been well known that, in contrast to mammalian cells, in trypanosomes the flagellar pocket membrane plays a pivotal role in protein trafficking between the interior and the exterior of the cell (27). Our findings elaborate the idea of the protease being transported outside the cell. Previous studies have also indicated the localization and exocytosis of a serine protease in Leishmania amazonensis through the flagellar pocket (31). The cellular location could be implicated in determining the physiological role of the protease and might be taken into account for future drug design strategies, although the exact mechanism of vesicle fusion, which might be involved in the transport process, still remains uncertain, and more-detailed study of the nature of this compartment needs to be performed.
Effect of antisera on infectivity for macrophages. We examined the effect of the anti-115-kDa extracellular protease antibody on in vitro invasion of macrophages. Early-passage promastigotes were incubated with the antibody against the 115-kDa protease. Promastigote internalization was inhibited by treatment of the L. donovani promastigotes with the antibody (Fig. 5). Microscopic analysis showed the conspicuous presence of numerous intracellular parasites in the host macrophages when promastigotes were not treated with the antibody, but the numbers of antibody-treated parasites in macrophages were significantly reduced (Fig. 5A). Thus, one can speculate that the parasites did not mediate internalization when treated with the antibody. Figure 5B shows that the numbers of invading parasites diminished after incubation of the antibody-treated parasites with macrophages for different times (up to 72 h), whereas the number of invading control parasites (either left untreated or treated with preimmune serum) showed successive increases as the incubation time increased. We might speculate that the antibody interacts with the extracellular protease and that this interaction somehow interferes with the invasion of macrophages by parasites. Pretreatment of macrophages with the purified 115-kDa enzyme prior to invasion permitted efficient invasion by parasites treated with the anti-115-kDa antibody (Fig. 5B). Importantly, the purified protease treatment appeared to be efficient in the restoration of parasite invasion. These results indicate the involvement of the protease in target cell invasion.
In vivo assay of L. donovani infectivity after immunization with the protease. To evaluate the efficacy of the 115-kDa serine protease as a potential protective antigen, female BALB/c mice were vaccinated with the protease together with Freund's complete adjuvant, subcutaneously or intraperitoneally, and were boosted three times at 2-week intervals. Four weeks after the final injection, the mice were challenged with 5 ϫ 10 5 or 5 ϫ 10 8 virulent L. donovani promastigotes. The mice were sacrificed 1 month following parasite challenge, and the parasite burdens in spleens and livers were estimated by counting the parasite numbers in those tissues. Mice immunized with the 115-kDa protease exhibited significant protection, as observed by dramatically lower parasite burdens than those for mice receiving Freund's complete adjuvant alone.
Mice challenged with 5 ϫ 10 5 parasites showed no discernible infection in either the spleen or the liver for as long as 8 months, but infection developed after 4 months in mice challenged with 5 ϫ 10 8 parasites (Fig. 6). Thus, vaccination against this protease is partially effective in eliciting a protective immune response. The mechanism of protection is currently unknown, and work is in progress to find out the basis of this protection. It could be due to an increase in the Th1/Th2 ratio, since several workers have reported that protective immunity in response to vaccination is associated with an increased Th1 response (12,28). Immunomodulatory cytokines might also be involved in eliciting the immune protection (12,28).
The localization of the 115-kDa protease inside the flagellar pocket, a specialized region of the plasma membrane involved in endocytosis and exocytosis, strongly suggests that this protease can be released into the extracellular medium by utilizing this vesicular compartment as the secretory pathway. On the other hand, the evidence that the 115-kDa secreted protease has strong proteolytic activity against extracellular matrix proteins, such as collagen and fibronectin, as demonstrated in our previous work (8), suggests that the protease is a superior agent for host tissue invasion. Our experimental data revealed that the relative abundance of the extracellular serine protease is greater in early-passage promastigotes than in late-passage and attenuated promastigotes. Therefore, this differential expression might play a crucial role in the life cycle of Leishmania donovani, and the molecular mechanisms that regulate the appearance of this protease in virulent, avirulent, and attenuated states should be further investigated. Previous reports have indicated that virulence for different Leishmania species is associated with different parameters, such as the length of in vitro culture (16), stationary versus log growth phase (17), and the expression of the major surface glycoprotein (34). We would infer from our findings that relative expression of the 115-kDa protease should be included in this list.
In summary, our work suggests the intracellular distribution of a novel type of serine protease, capable of degrading the ECM, in the flagellar pocket region of L. donovani. The expression of this protease at relatively high levels in early-passage promastigotes compared to levels in late-passage and attenuated stages suggests that the 115-kDa serine protease could contribute to parasite invasion and virulence. Taken together, these observations could help us to acquire important knowledge about the mode of action of this protease, at least in this particular species of Leishmania, and this protease can be considered as a potential target for antileishmanial drug design, to help eradicate this devastating disease affecting millions around the world.

ACKNOWLEDGMENTS
Thanks are due to Amritlal Mandal (Department of Physiology, University of Arizona, Tucson, AZ) and S. Roy (Infectious Diseases and Immunology Division, IICB, Kolkata, India) for help with our research.
Financial assistance from the Indian Council of Medical Research (New Delhi, India) and the Council of Scientific and Industrial Research (New Delhi, India) is acknowledged.


extfieldsofstudy: Biology Medicine

s2fieldsofstudy: Biology Medicine

text:
Downregulation of both gene expression and activity of Hsp27 improved maturation of mouse oocyte in vitro

Background Heat shock protein 27 (Hsp27), a member of the small heat shock protein family, is an apoptosis regulator. Our previous proteomic study showed that Hsp27 mainly expressed in human oocyte, and that Hsp27 expression was downregulated in the ovaries derived from women with the polycystic ovary syndrome (PCOS), a well known endocrinal disorder with abnormal apoptotic activity and folliculogenesis. However, the exact effects of Hsp27 downregulation on oocyte development have not yet been clarified. Methods The expression of Hsp27 gene was downregulated in the mouse oocytes cultured in vitro using siRNA adenovirus infection, while the activity of Hsp27 was decreased by microinjection of polyclonal Hsp27 antibody into the cytoplasm of germinal vesicle (GV) oocytes. Oocyte maturation rate was evaluated by morphological observation. Early stage of apoptosis was determined using Annexin-V staining analysis and some critical apoptotic factors and cytokines were also monitored at both mRNA level by real time RT-PCR and protein expression level by immunofluorescence and western blot. Results Hsp27 expressed at high level in maturing oocytes. Infection with AdshHsp27, and microinjection of Hsp27 antibody into GV oocytes, resulted in the improved oocyte development and maturation. Germinal vesicle breakdown (GVBD) rates were significantly increased in two AdshHsp27-treated groups (88.7%, 86.0%) and Hsp27 antibody-injected group (77.0%) when compared with control (76.2% in AdGFP, 64.4% in IgG-injected), respectively. In addition, the rates of metaphase II (MII) development in two AdshHsp27-treated groups (73.8%, 76.4%) and Hsp27 antibody-injected group (67.3%) were higher than that in the controls (59.6% in AdGFP, 55.1% in IgG-injected). We also found that the rates of early stage of apoptosis in Hsp27 downregulated groups (46.5% and 45.6%) were higher than that in control group (34.1%) after 8 h of IVM. Similarly, downregulation of Hsp27 caused a significantly enhanced the expression of apoptotic factors (caspase 8, caspase 3) and cytokines (bmp 15 and gdf 9). Conclusions Downregulation of Hsp27 improved the maturation of mouse oocytes, while increased early stage of apoptosis in oocytes by inducing the activation of extrinsic, caspase 8-mediated pathway.

In the ovary, apoptosis has been implicated in the granulose cells of atretic antral follicles and in regressing corpora lutea [18][19][20][21][22]. Derangement of apoptotic activity was observed in PCOS ovary tissue with the altered expres-sion of apoptotic-related regulators, including heat shock proteins (Hsp 90, Hsp 10), nuclear receptor subfamily, dickkopf homologue 3, and so on [23][24][25][26]. Hsp27, belonging to the small heat shock protein family, is a molecular chaperone protein involved in cellular protection in response to a variety of stresses such as heat shock, toxicants, injury, and oxidative stress [27][28][29][30]. Emerging evidences show that Hsp27 has strong anti-apoptotic properties by interacting directly with the caspase activation components in apoptotic pathways, consequently exerting protective effects in apoptosis-related injuries [31][32][33][34]. Interestingly, our previous proteomic study showed that Hsp27, a strong anti-apoptotic regulator, mainly localized in human oocyte, and was downregulated in the ovaries derived from women with PCOS [35]. However, the alteration of apoptotic activity, as well as effect of Hsp27, in PCOS ovaries needs to be further clarified.
Our hypothesis was that Hsp27 and its related pathways could have some effects on oocyte development, maturation, apoptosis and cell cycle in vivo and in vitro, and even participate in the follicle development and PCOS pathophysiology. In this pilot study, we firstly investigated the effect of Hsp27 downregulation on the meiotic progression and apoptosis in mouse oocyte model cultured in vitro.

Animals
The ICR mice were fed ad libitum with a standard diet and maintained in a temperature and light-controlled room (20-22°C, 12/12 h light/dark), in accordance with the Animal Research Committee Guidelines of Nanjing Medical University.

Collection and culture of mouse oocytes
Germinal vesicle (GV) oocytes were collected from 6week-old female ICR mice. 46-48 h previously, mice were received an intraperitoneal injection of 10 IU of pregnant mare serum gonadotropin (PMSG, Folligon, Intervet, Castle Hill, Australia). Mice were sacrificed, and ovaries were placed in M2 medium (Sigma, St. Louis, MO). Cumulus oocyte complexes were recovered from ovaries by repeatedly puncturing the surface with fine steel needles, and cumulus cells were removed by hyaluronidase treatment (Sigma, 300 U/ml in PBS) under a dissecting microscope [36]. For preparation of zona pellucida-free oocytes, the oocytes were then exposed to acidic Tyrode's solution (pH 2.5-3.0) with aspiration of the oocyte in and out of a glass micropipette to remove the zona pellucida [37]. Usually the zona pellucida was only partially dissolved and it could be removed by the pipette within 30s. Ding et al reported that zona-free oocytes didn't affect normal fertilization and blastocyst development [38].
After being immediately transferred to M2 and washed three times, 20-25 zona-free oocytes per group were put into 32 μl droplets of maturation medium of M16 (Sigma) containing 10% fetal bovine serum (FBS; Invitrogen, Grand island, NY) in a 5% humidified atmosphere at 37°C under a layer of mineral oil. These prepared zona-free oocytes were then used in either control or treatment groups for further investigation.
Zona-free oocytes were incubated with adenovirus at the same multiplicity of infection (MOI) in medium at 37°C for specific durations as indicated in the following experiments.

Microinjection of Hsp27 antibody into GV oocytes
Hsp27 antibody (stock solution, 200 ug/ml, goat polyclonal, Santa Cruz Biotechnology, Santa Cruz, CA) was microinjected into the cytoplasm of GV zona-intact denuded oocytes as described by Dai et al [44]. Goat normal IgG-injected (Santa Cruz, CA) oocyte and untreated oocytes were used as controls to assess injection itself damage. To determine maturation rate in vitro, the microinjected GV oocytes were cultured in M16 medium (Sigma) in a 5% CO 2 incubator at 37°C for 14 h. Oocytes without GVs were scored as germinal vesicle breakdown (GVBD) stage, while those with a polar body scored as metaphase II (MII). The microinjection experiments were done 10-12 replicates, using a total of > 300 oocytes. A Nikon Diaphot ECLIPSE TE 300 inverting microscope (Nikon UK Ltd, Kingston upon Thames, Surrey, UK) equipped with Narishige MM0-202N hydraulic threedimensional micromanipulators (Narishige Inc., Sea Cliff, NY) was used. Microinjection was completed in 40 min, with volume of about 5-7 pl per oocyte.

Annexin-V staining in oocytes
Staining was performed with an Annexin-V kit according to the manufacturer's instructions (KenGentec, Nanjing, China). Annexin-V, a phospholipid-binding protein, detects the translocation of phospholipid phosphatidylserine (PS) from the inner to the outer cytoplasmic membrane, which is known to occur during the early stages of apoptosis. At the same time, samples were also stained with propidium iodide (PI) to distinguish live cells from dead cells. Briefly, zona pellucida-free oocytes infected with siRNA adenovirus via co-culturing for 1.5 h and 8 h were washed twice in PBS and stained with 500 μl of binding buffer, which contained 5 μl Annexin-V-fluorescein isothiocyanate (FITC) and 5 μl PI for 5-15 min in the dark. Following this, samples were mounted on siliconized slides and observed under a laser confocal scanning microscope (ZEISS Fluorescent Microsystems, Göttingen, Germany).

Real time RT-PCR analysis
To determine mRNA abundance, real-time RT-PCR analysis was performed using ABI 7300 (Applied Biosystems, Foster City, CA). RNA isolation was accomplished using the RNeasy Micro Kit (Qiagen, Valencia, CA) from the pooled oocytes (20-25 oocytes/tube). In vitro reverse transcription was carried out using Sensiscript Reverse Transcription Kit (Qiagen) with oligo-dT primer at 37°C for 60 minutes. For the real time RT-PCR reaction, cDNAs were used as templates for amplification to quantify the steady-state mRNA levels of the tested genes using Quanti Tect SYBR Green PCR Kits (Takara Shuzo Co Ltd, Kyoto, Japan). Relative quantitation of target gene expression was evaluated by the 2(-Delta Delta Ct) method [45], and the experiment was repeated at least three times using different sets of oocytes. In our study, beta-actin and GAPDH gene were used as the internal standards referred to similar papers [46,47]. To ensure only target gene sequence-specific, non-genomic products were amplified by real-time RT-PCR, careful design and validation of each primer pair. Primers used for real time RT-PCR are shown in Table 1.

Immunofluorescence
Oocytes of different treatment groups were fixed in 4% paraformaldehyde in PBS (pH7.4) for at least 30 min at room temperature and then incubated in permeabilization buffer (0.5% Triton X-100 in 20 mM Hepes 3 mM MgCl 2 , 50 mM NaCl, 300 mM sucrose, and 0.02% NaN 3 )   The membrane was then washed 3 times in TBS-Tween (TBST) for 10 min each, and processed using an enhanced chemiluminescence detection system (Alpha Innotech, San Leandro, CA). The molecular weights of the detected proteins were deduced by comparison with recombinant molecular weight standards (New England BioLabs, Ipswich, MA).

Statistical analysis
Each experiment was repeated at least three times. All data are presented as the mean ± SD, and one-way analysis of variance and a log linear model were used to compare the mRNA and protein levels. Chi-square analysis was used to compare the rates of oocyte maturation and early stage of apoptosis. A value of P < 0.05 was considered statistically significant, and if P < 0.01, it was noted.

Construction of recombinant adenovirus Ad-shHsp27 and its expression
AD-293 cells were infected by the adenoviruses packaged with hairpin siRNA targeted against Hsp27/control. The results showed that the suppression rate of Hsp27 expression was as high as 75% vs. control (Data not shown, P < 0.05) after infecting with Ad-shHsp27 and control for 48 h.
To examine the reduction degree of Hsp27 mRNA in mouse oocytes after infected with Ad-shHsp27 adenovirus, real time RT-PCR was performed. As Figure 1B displayed, Ad-shHsp27 significantly reduced endogenous Hsp27 mRNA abundance in mouse oocytes (50%). The suppression degree of Hsp27 at protein level was detected by immunofluorescence ( Figure 1A), which was in accor-dance with the real time PCR. These results suggested that the knockdown of Hsp27 expression by Ad-shHsp27 infecting was effective. Therefore, these two siRNA were chosen for the subsequent experiments.

Localization and expression of Hsp27 in the maturing oocytes
The localization of Hsp27 protein in maturing mouse oocytes was showed in Figure 2A. Hsp27 expression was recognized mainly in the cytoplasm and nuclei (except the nucleolus) of mouse oocytes. To assess the levels of Hsp27 mRNA in the maturing oocyte, real time RT-PCR analysis was performed using cDNAs equivalent in oocytes at different developmental stages ( Figure. 2B). Hsp27 expression dramatically increased following oocyte maturation, Western blot analysis confirmed that Hsp27 at protein level also increased following oocyte maturation, and dramatically increased in MII ( Figure. 2C), which was consistent with the results of real time RT-PCR and immunofluorescence.

Microinjection of Hsp27 antibody into mouse oocytes
Polyclonal antibody of Hsp27 was microinjected into oocytes cytoplasm at GV stage to downregulate the activity of Hsp27. After 3 h of microinjection, the GVBD rate (77.0%) significantly increased, when compared with normal control (67.3%) or IgG-injected control (64.4%) (P < 0.01, vs IgG-injected group). After 14 h of microinjection, the maturation rate of MII stage (67.3%) was also higher than those of two controls (below 60%) (P < 0.01, vs IgGinjected group; Table 2). These findings suggested that the lowed Hsp27 activity in mouse oocytes could improve the maturation of oocytes.

Annexin-V staining of oocytes
Oocytes were classified into three groups as Anguita et al described [48]. The first group contained necrotic oocytes with PI-positive red nuclei, indicative of membrane damage. Oocytes with a discontinuous green signal originating from the remnant portions of the membrane were viable non-apoptotic oocytes (Figure. 3A). The second group contained viable oocytes which were negative for Annexin-V staining (Figure. 3B). The third group con-sisted of early apoptotic oocytes with homogeneous Annexin-V-positive signal in the membrane (Figure. 3C).
The ratios of Annexin-V-positive cells in two shHsp27 groups were higher than negative control (Table 4). After 3 h of treatment, the early apoptotic rate of oocytes infected with Ad-shHsp27 (38.9%, 36.2%) was not significantly different from control (33.8%). The ratios of early apoptotic oocytes in shHsp27-treated groups after 8 h of treatment were 46.5% and 45.6%, which were significantly higher than the negative control (34.1%). The results suggested that Hsp27 downregulation in oocytes could promote the early stage of apoptosis.

Variation of downstream apoptosis-related factors and oocyte secreted factors with defect of Hsp27 protein
To elucidate the involved apoptotic pathway(s), the expressions of four apoptotic factors (caspases 8, caspase 9, caspase 3 and cytochrome c) were measured in oocytes following different treatment by real time RT-PCR, immunofluorescence and western blot. As shown in Figure 4 and Figure 5, the results of these assays were accordant. Hsp27 downregulation in mouse oocytes dramatically increased the activitied caspase 8 and cas- Table 4: The rate of early stage of apoptosis in oocytes at 3 and 8 hours after siRNA adneovirus infection, was evaluated by Annexin-V staining in different groups (Control, shHsp27(1), shHsp27(2)).  pase 3, while caspase 9 and cytochrome c activities did not change. These results suggested that downregulation of Hsp27 in mouse oocytes resulted in activation exclusively of the extrinsic, caspase 8-mediated apoptotic pathway, rather than the intrinsic, caspase 9-mediated pathway.
In addition, the expressions of two important oocytederived growth factors (bmp 15 and gdf 9) after downregulation of Hsp27 in mouse oocytes were also investigated ( Figure. 4A and 4B). The results indicated that bmp 15 and gdf 9 were increased significantly in oocyte after downregulation of Hsp27.

Discussion
In the present study, the effect of Hsp27 downregulation on oocyte development and maturation was investigated in the mouse oocyte model cultured in vitro. Expression of Hsp27 gene was downregulated in oocytes using siRNA adenovirus infection, while the activity of Hsp27 was decreased by microinjection of polyclonal Hsp27 antibody into mouse GV oocytes. Interestingly, our results showed that Hsp27 downregulation in mouse oocytes improved oocyte maturation and increased oocyte early stage of apoptosis.
Hsp27 was investigated as an antiapoptotic factor [49][50][51]. Hsp27 could directly bind and co-precipitate with cytochrome c, consequently leading to activation of the caspase 9 and caspase 3 in mouse, rat and human cells [31,33,52,53]. Kamradt et al reported that Hsp27 could negatively regulate cytochrome c-and caspase 8-dependent activation of caspase 3 in human breast carcinoma cells [54]. To our knowledge, there were few reports about the roles of Hsp27 in oocyte. In this pilot study, after downregulation of Hsp27 in mouse oocytes, the ratio of early stage of apoptosis was significantly increased. Meanwhile, the expression of major apoptotic factors i.e. caspase 3, caspase 8 was dramatically increased. These results suggested that downregulation of Hsp27 in mouse oocytes might increase early stage of apoptosis by inducing the activation of the extrinsic, caspase 8-mediated apoptotic pathway.
Our results showed that the lowed levels of Hsp27 significantly increased GVBD rate and MII rates in both siRNA infection group and antibody microinjection group when compared with controls, indicating that Hsp27 downregulation was positively related to oocyte maturation. This result was also supported by detecting the expression of two important oocyte-derived growth factors-bmp 15 and gdf 9, which were known to be responsible for controlling fundamental physiological processes in oocyte development and follicular growth [55]. Juengel et al reported that immunization against gdf 9 and bmp 15 alone or together in cattle oocytes could block folliculogenesis and reduce follicular size, which indicated the critical role of bmp 15 and gdf 9 on oocyte development and follicular growth [56]. In this study, we found that bmp15 and gdf 9 were noticeably upregulated after Hsp27 downregulation in the mouse oocytes model cultured in vitro, suggesting that there was an enhanced effect of downregulated Hsp27 on oocyte development.
Recently, there were numerous studies focused on the relation between early stage of apoptosis and oocyte developmental competency in the pooled oocytes cultured in vitro [12,48,57,58]. As we known, early stage of apoptosis was regarded as a sequential, but reversible, process of cell death [10,57]. As Anguita et al and Jaroudi et al indicated, the oocytes undergoing early stage of apoptosis did not mean that they must develop into late apoptosis, early stage of apoptosis oppositely decreased their developmental competence [48,59]. Bilodeau-Goeseels and Panich proved that oocytes with early signs of atresia had good developmental potential by detecting the transcriptional activity in early bovine embryos from different classes of cumulus-oocyte complexes [60]. Additionally, some clinical studies provided consistent results that oocytes from slightly atretic COCs with signs of cumulus expansion had a better embryonic developmental capacity after IVF than those considered to be of the highest quality [61][62][63][64][65]. Concordantly, the percentage of apoptotic cumulus cells increased when COCs were exposed to FSH. Oocytes from those COCs exhibited a higher developmental potential in terms of blastocyst formation rate [66][67][68]. Those reports suggested that early stage of apoptosis had a positive relation with the improved oocyte developmental potential. In our present study, downregulation of Hsp27 led to a lower antiapoptotic effect and a significant increase in the GVBD rate and MII rate, which was consistent with previous reports [57]. Taking these results together, we speculated that downregulation of Hsp27 induced early oocyte apoptosis, consequently improving oocyte development. In fact, it is difficult at present to understand the molecular mechanism of the correlation between early stage of apoptosis and subsequent maturation of oocyte in follicles in PCOS. Our hypothesis is that downregulation of Hsp27 could be one of the bridges between those events, which is our objectives in future studies.
Previous proteomic analysis comparing normal and PCOS ovarian protein profiles identified that expression of Hsp27 in PCOS ovaries was decreased when compared with normal ovarian tissues [35]. PCOS is known as the most common cause of anovulatory infertility, resulting from a disorder of follicular maturation of uncertain aetiology [15]. Several reports showed that apoptosis was perturbed in PCOS ovaries, with overexpression of apoptotic factors and downregulation of anti-apoptosis factors. However, the others showed the reversed results [69][70][71]. Das et al reported that there were significant differences in cell death rate and proliferation rate of granulosa cell populations in PCOS patients [72,73]. Oocyte development is interdependent with its surrounding granulose cells [11], and disfunction of granulose cells may contribute to the abnormal folliculogenesis observed in PCOS [5,74,75]. Interestingly, we found in this study that the downregulated Hsp27 improved oocyte maturation, which seems to be contradicted with the atresia follicles observed in PCOS. One possible explanation for this contradiction could be that our research was performed in mouse oocyte model cultured in vitro. This model simulates the normal oocyte's development, which could be different from the pathological procedure of PCOS characterized by an abundance of large but immature follicles. We need more studies on the effects of Hsp27 in PCOS pathophysiology in future studies.

Conclusions
In conclusion, evidence provided in the present study indicated that downregulation of Hsp27 improved oocyte maturation, and increased early apoptosis of mouse oocytes by triggering the extrinsic, capsase 8-mediated pathway. This is the first report, to our knowledge, to establish a direct link between Hsp27 and oocyte maturation. Relative deficiency of Hsp27 expression in GV oocytes may contribute to the disordered oocyte development in PCOS. Further studies will be required to elucidate the role of Hsp27 in oocyte maturation in vivo and PCOS pathophysiology. Oocytes were fixed in 4% paraformaldehyde after microinjecting of Hsp27 antibody for 36-48 h. Then these oocytes were stained with cleaved-caspase 3 antibody (green). Chromosome material was counterstained with propidium iodide (red). A) IgG-injected oocyte. B) Hsp27 antibody-injected oocyte.


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Colorimetric Immunoassay for Detection of Tumor Markers

Tumor markers are substances, usually proteins, produced by the body in response to cancer growth, or by the cancer tissue itself. They can be detected in blood, urine, or tissue samples, and the discovery and detection of tumor markers may provide earlier diagnosis of cancer and improved therapeutic intervention. Colorimetric immunoassays for tumor marker detection have attracted considerable attention, due to their simplicity and high efficiency. The traditionally used colorimetric immunoassays for the detection of tumor markers are based on enzyme-linked immunosorbent assays, and the great achievement of nanotechnology has further opened opportunities for the development of such kind of immunoassays. This paper will summarize recent advances in the field of colorimetric immunoassays for detecting tumor markers, which is aimed to provide an overview in this field, as well as experimental guidance for the learner.

Introduction
Cancer is a leading cause of death worldwide, causing about 13% of all human deaths in 2007 (7.6 million), and deaths from cancer are projected to continue rising, with an estimated 12 million deaths projected for 2030. According to the World Health Organization, more than 30% of cancer deaths could be prevented if the cases were detected early and treated [1]. So, one key point of ultimate success for cancer treatment is the accurate diagnosis at the early stages.
Traditional methods for the diagnosis of cancer are endoscopy, biopsy and cytology specimen tests, as well as imaging/radiology tests, such as X-ray, positron emission computed tomography-computer tomography (PET-CT) and magnetic resonance imaging (MRI) [2][3][4][5]. Unfortunately, these diagnostic methods are not very powerful for the diagnosis of cancer at early stages, and some of these methods are quite time-consuming, expensive; thus not available for to a large number of people [6]. Therefore, the development of simple and rapid strategies that are specific and reliable for the diagnosis of cancer at early stages is of utter importance.
It has been known that cancer cells or other related non-tumor cells can release specific tumor markers, which are usually proteins, as tumors develop, into the circulation system in response to cancer growth. These tumor markers can be detected in blood, urine, or tissue samples, and the level of them is associated with the stage of cancer [7][8][9][10]. Alongside the development of proteomic technologies, lots of protein tumor markers have been discovered for many types of cancer (Table 1) [11][12][13][14][15][16][17][18]. Considering the high importance and potential application in the early diagnosis of cancer, detection of tumor markers has received more and more attention [19][20][21][22][23]. While some novel immunoassays have been developed [24][25][26][27][28][29][30][31][32][33], a variety of techniques have been employed for the development of these assays. Table 2 summarizes some examples of immunoassays which have been developed for tumor markers analysis [33][34][35][36][37][38][39][40][41][42][43][44][45][46][47][48][49]. Among the various detection techniques, the colorimetric method, in which the event is disclosed through a visual color change in the reaction medium, has proven the most convenient [50,51]. Thus, remarkable progress has been made on the design of colorimetric sensing systems for tumor marker detection over the years. Table 1. Some tumor markers currently in use.

Tumor Markers Related Cancers Usual Sample
Refs.
Among the nanomaterials employed for the colorimetric detection of tumor markers, Au-NPs are the most commonly used, due to their unique optical properties. Their optical properties are strongly dependent on not only the size but also aggregation state of the particles, so smaller individual nanoparticles appear wine red while larger particles or aggregates of smaller particles range from purple to deep blue [45]. Besides, their excellent biocompatibility makes Au-NPs able to be fashioned with a wide range of biomacromolecules such as nuclear acids and enzyme-linked antibodies, greatly extending their application in the colorimetric detection of tumor markers. Therefore, large numbers of Au-NPs based colorimetric immunoassays have been developed for the detection of tumor markers, oncogenes and even tumor cells [66][67][68][69][70].
Apart from Au-NPs, magnetic particles (MPs) represent another exciting prospect in current analytical fields because they can be easily separated from a matrix by using a magnetic field [71][72][73]. Taking advantage of their unique magnetic characteristic as well as excellent biocompatibility, MPs have been widely utilized as a universal separation tool in the fabrication of colorimetric sensing systems for the detection of tumor markers [38,44,45,49,65,70].
Blocking buffer: PBS containing 0.1% bovine serum albumin (BSA) and 0.02% thimerosal. All buffer reagents and other inorganic chemicals can be supplied by Sigma, Aldrich, or Fluka. All chemicals are used as received, and all aqueous solutions are prepared with doubly distilled water.

Nanomaterial-Based Colorimetric Immunoassays
Monodispersed Au-NPs with size from 2 to 250 nm can be commercially received from Ted Pella Inc. (

ELISA-Based Colorimetric Immunoassays
The most frequently used ELISA method in colorimetric immunoassays for tumor marker detection is the two antibody -sandwich‖ ELISA, which measures the amount of antigen between two layers of antibodies [52]. In this assay, a capture antibody is firstly immobilized onto a solid support. The tumor marker, which is present in a biological sample or standard mixture, is then bounded and concentrated onto the support surface during incubation. After that, the solid support is incubated with a solution containing the detection antibody, thus the detection antibody will also bind with the tumor marker. Since the detection antibody is also tagged with certain enzymes, the color change caused by the enzyme-catalytic reaction can be thus used to quantify the amount of target tumor marker. The experimental details are described below, mainly with a slightly modified procedure recommended by Abcam [74].

Coating with Capture Antibody
Coat the wells of a microtiter plate with the capture antibody at a concentration of 1-10 μg/mL in carbonate/bicarbonate coating buffer. Then cover the microtiter plate with an adhesive plastic. After incubation overnight at 4 °C, remove the coating solution and wash the plate at least twice by filling the wells with 200 μL of washing buffer. The solution in the wells can be removed by flicking the plate over a sink, and the remaining drops in the wells can be removed by patting the plate on a paper towel.

Blocking and Adding Samples
200 μL of blocking buffer containing 5% non fat dry milk is firstly added into each well to block the remaining protein-binding sites. After that, the plate is covered with an adhesive plastic and incubated with the blocking buffer for at least 1-2 h at room temperature or overnight at 4 °C. Then add 100 μL of appropriately diluted samples into each well, and incubate the plate with the sample solution for 90 min at 37 °C. After incubation, remove the samples and wash the plate twice by filling the wells with 200 μL of washing buffer.

Incubation with Detection Antibody
Dilute the detection antibody appropriately with 0.1 M bicarbonate buffer (pH 9.2). Then, add 100 µL of the buffer solution to each well. Cover the plate with an adhesive plastic and incubate for 2 h at room temperature. After that, wash the plate four times with washing buffer.

Colorimetric Detection
Although many different types of enzymes have been used in ELISA, HRP and ALP are the two most widely used enzymes [43]. The substrate for HRP is H 2 O 2 , and the cleavage of H 2 O 2 is usually coupled with the oxidation of a hydrogen donor which may change the color of the test solution during the reaction, so hydrogen donors should also be involved in the reaction. These hydrogen donors, usually TMB, OPD and ABTS, are light sensitive, so they need to be stored in the dark. Furthermore, OPD is considered hazardous (potential carcinogens), so it always needs to be handled with care. In a typical experiment, H 2 O 2 and hydrogen donors are added into each well and incubated for 15-30 min. After that, equal volume of stopping solution (2 M H 2 SO 4 ) is added to stop the reaction. Finally, the absorbance of the product of TMB oxidation is measured at the wavelength of 450 nm, while OPD at 492 nm; ABTS at 416 nm.
The most widely used ALP substrate is pNPP. The yellow color of the product nitrophenol can be measured at 405 nm after 15-30 min incubation at room temperature, and this reaction can be stopped by adding equal volume of 0.75 M NaOH.

Nanomaterial-based Colorimetric Immunoassays
The unique properties of nanomaterials may offer a wide range of opportunities in colorimetric immunoassays of tumor markers. In the following section, we provide experimental details for the synthesis, surface modification and utilization of nanomaterials in the fabrication of colorimetric sensing systems for the detection of tumor markers.

Synthesis of 13 nm Au-NPs
All glassware used in the synthesis procedures should be firstly immersed in freshly prepared aqua regia (HNO 3 : HCl = 1:3) for 30 min, then washed with water and dried before use. Au-NPs are synthesized by reducing HAuCl 4 with Citrate· 3Na. Briefly, a 100 mL aqueous solution of 0.01% (w/v) HAuCl 4 is added into a round-bottom flask and stirred to boil. Then, 3.5 mL 1% trisodium citrate is added rapidly into the boiling solution, the color of which changes from colorless to wine red after boiling for another 15 min with vigorous stirring. The size of the nanoparticles is 12.5 ± 2.3 nm, which can be determined by transmission electron microscope (TEM). The concentration of Au-NPs is 3.5 nM, which can be calculated from the quantity of starting material (HAuCl 4 ) and the size of Au-NPs at a wavelength of 520 nm [45].

Modification of Au-NPs
The main technique reported for the modification of Au-NPs surfaces is based on Au-S covalent bond formation between the modified molecules and the gold atoms on the particle surface [71], which makes use of sulphur-containing ligands, such as thiol, disulphide and thiolester.

Modification of Au-NPs with Thiol-oligonucleotides
The above prepared Au-NPs solution is firstly condensed by centrifugation at 12,000 rpm for 20 min at 4 °C to a 1 mL Au-NPs solution, while 800 μL supernatant is removed; the remaining 200 μL is redispersed. After condensation, 800 μL Au-NPs solution is mixed with 200 μL, 5 μM 5'-thiol-oligonucleotides at room temperature. After 24 h duration in darkness, the mixture is centrifuged twice, each for 20 min at 12,000 rpm to remove the excess thiol-oligonucleotides. After centrifugation, the precipitate is washed with 4 mM Citrate· 3Na. The Au-NPs, modified with thiol-oligonucleotides, are stored at 4 °C when not in use. To determine the number of thiol-oligonucleotides modified on each Au-NP, 2-mercaptoethanol (1.0 M, 10 μL), which has a smaller size than oligonucleotides and is easier to be modified on the surface of Au-NPs through Au-S bond, is used to replace thiol-oligonucleotides from the surface of Au-NPs (11 nM, 990 μL). The amount of replaced thiol-oligonucleotides in the supernatant liquor can be thus used to calculate the quantity on each Au-NP [75].

Modification of Au-NPs with Antibodies
The methods to modify Au-NPs with antibody include adsorption [76], N-hydroxysuccinimide (NHS) ester chemistry [77], and oligonucleotide-directed immobilization [78]. Among them, the direct adsorption method is the simplest one, due to the fact that neutral or weakly negative-charged proteins can be adsorbed on the surface of Au-NPs easily and stably through coordinate-covalent bond [79]. The preparation of Au-NPs modified with antibodies through direct adsorption is as follows [45]: 20 μL of protein solution (1 mg/mL) is first added into 5 mL of the above prepared Au-NPs solution, and the pH is appropriately adjusted by adding 1 M NaOH solution. Then the mixed solution is incubated for 30 min at 37 °C with gentle shaking, and centrifuged to 1 mL (12,000-13,000 rpm, 20-30 min, 4 °C). After that, 100 μL of 10% (w/v) BSA solution, used as non-specific blocker, is added into the solution to passivate the surface of Au-NPs. After 30 min, the mixed solution is again centrifuged (12,000-13,000 rpm, 20-30 min, 4 °C ). The supernatant is subsequently decanted, and the precipitate is rinsed with 1 mL of PBS buffer. The centrifuging/rinsing procedure should be repeated three times to remove the unbounded antibodies and BSA.

Modification of MPs
MPs may provide an efficient tool of separating target analytes from the liquid suspension; however, unmodified MPs have some limitations in the applications. Therefore, MPs surfaces should first of all be modified with some specific linking groups for further biomolecules binding, including: (1) Active chemical groups (e.g., carboxyl and amino groups), which can covalently bind biomolecules in the presence of specific cross-linking reagents; (2) streptavidin/avidin ligands, which can be specifically attached to biotinylated biomolecules [71][72][73]. Taking into account that amino-or streptavidinfunctionalized MPs are commercially available, the following is only a description of the modification by antibodies for these pre-functionalized MPs.

Modification of Amino-Functionalized MPs
Amino-functionalized MPs can be modified with antibodies by using glutaraldehyde-amine coupling chemistry [33]. Briefly, amino-functionalized MPs (0.05 mM) in 1 mg mL -1 of ethylenediaminetetraacetic acid (EDTA) solution (5 mL) are firstly washed with 5% (v/v) pyridine washing buffer (10 mL). Then, MPs are separated magnetically. This process should be repeated three times. After that, the MPs are activated by glutaraldehyde in pyridine washing buffer (5 mL, 5%) for 3 h at room temperature, separated magnetically twice, and resuspended in pyridine washing buffer (10 mL). Finally, antibody dissolved in pyridine wash buffer (1 mL at 1 mg mL -1 ) is added to the MPs solution. After 10 h duration at room temperature, 1 mg BSA is added to the MPs solution. After another 10 h at room temperature, the MPs are washed twice and then resuspended in pyridine washing buffer (5 mL). Subsequently, glycine solution (3 mL, 1 M at pH 8.0) is added into the above solution to quench all of the unreacted aldehyde groups, and the resulting solution is stirred for 30 min. After magnetic separation three times and the washing step, MPs modified with antibody (Ab-MPs) are resuspended in PBS solution.

Modification of Streptavidin-Functionalized MPs
Streptavidin-functionalized MPs are firstly washed three times with washing buffer (75 mM NaCl, 7.5 mM Citrate· 3Na, pH 7.0) prior to use. Then they are diluted to 0.1 mg mL -1 with PBS, pH 7.4. After that, a solution of biotinylated antibody is added to the MPs solution at a ratio of 100 μg protein to 1 mg MPs, and the mixed solution is incubated for 20 min at room temperature. Finally, the Ab-MPs are thoroughly washed three times with pH 7.4 PBS [45].

Magnetic Collection and Separation of Target Tumor Markers
MPs modified with capture antibodies can be used for fast target protein collection and separation. In a typical experiment, 50 µL of capture antibody-modified MPs (0.2 mg mL −1 ) are first added to a 200 µL eppendorf tube. Then, 50 µL of assay buffer containing various concentrations of target tumor markers are added. After incubation for 30 min at 37 °C with gentle shaking, the magnetic beads are magnetically collected and twice rinsed with washing buffer [70].

Application of Au-NPs in Colorimetric Immunoassays of Tumor Markers
Basically, two colorimetric procedures by making use of Au-NPs have been proposed for tumor marker quantification: (a) Analysis of the state change of Au-NPs in solution, including the homogenous growth and aggregation triggered by a biological process; (b) analysis of the activity of enzymes that attached to Au-NPs, in which case Au-NPs act as enzyme carriers and signal enhancers.

Colorimetric Assay Using Homogenous Growth of Au-NPs
10 mL solution for the growth of Au-NPs is first prepared in a 20 mL vial, consisting of final effective concentrations of 0.25 mM HAuCl 4 and 0.1 M cetyltrimethylammonium bromide (CTAB). Before use, the solution is gently mixed with ascorbic acid (AA) solution with a final AA concentration of 10 mM and stirred. Seperately, Au-NPs/tumor marker complexes are first separated through magnetic separation and washed with 0.01 M PBS buffer (pH 7.4) three times to remove the excess of free Au-NPs. Then, Au-NPs are dissociated from the complexes by using 50 mM NaCl/1 M NaOH elution solution. Finally, the upper aqueous solution containing the dissociated Au-NPs is transferred to a 1 mL cuvette, followed by addition of the above previously prepared 1 ml solution for the growth of Au-NPs. The growth of Au-NPs and the color change of the solution are then recorded [38].

Colorimetric Assay Using Aggregation of Au-NPs
In this section, we introduce the experimental details of two typical assays. One is based on single chain fragment variable recombinant antibody (scFv)-functionalized Au-NPs, while the other is colorimetric bio-barcode assay.

scFv-Functionalized Au-NPs Based Assay
Firstly, scFv-functionalized Au-NPs are prepared as described in Section 3.2.1.2.2. Then, a range of concentrations of target protein are added to the solution of the scFv-functionalized Au-NPs while stirring, thus the scFv-functionalized Au-NPs show visible color change due to the aggregation of the nanoparticles caused by the interaction of scFv with the target protein. After 30 min, the developing reaction can be recorded by using a UV-visible spectrophotometer [69].

Colorimetric Bio-Barcode Assay
In a typical experiment, two types of probes are prepared. The first is a magnetic probe, i.e., MP modified with monoclonal antibodies for target tumor marker, prepared as described in Section 3.2.1.3.1. The second is a barcode probe, i.e., a porous silica particle modified with the monoclonal antibodies and barcode DNA complements. To prepare the barcode probe, amino-functionalized porous silica microparticles are firstly linked to antibodies using glutaraldehyde-amine coupling chemistry. Then, 3'-amino-functionalized barcode DNA complements (1 mL at 100 µM) are added to the solution containing the antibody modified silica particles. After incubation overnight at room temperature with gentle shaking, the solution is centrifuged twice, and the pellets are resuspended in 1 mL of 0.2 M ethanolamine. After 30 min duration at room temperature, centrifugation is performed again to remove the supernatant, and a 10% BSA solution is subsequently added, followed by repeating the centrifugation steps twice. Finally, the resulting pellets are resuspended in 1 mL of 0.15 M PBS solution, which is further mixed with 30 µL 100 µM barcode DNA solution and incubated for 1 h at room temperature.
In the tumor marker detection assay, 15 µL of magnetic probe solution (1.5 × 10 9 beads/mL) is first added to 20 µL of tumor marker solution, followed by the addition of 15 µL of barcode probe solution (1 × 10 9 beads/mL). The mixed solution is incubated at 37 °C for 50 min on an orbital shaker. Then, the solution is magnetically separated as described in Section 3.2.2, and the magnetically separated complexes are washed with 0.15 M PBS solution three times. After that, 50 µL of NANOpure water (18 MΩ) is added to the complexes, and the solution is kept on a rocking shaker at 70 °C for 10 min to release the barcode DNA. Finally, the supernatant containing free barcode DNA strands is collected for barcode DNA detection. To detect the barcode DNA, DNA-functionalized Au-NPs (prepared as described in Section 3.2.1.2.1) are added to the barcode DNA in 0.15 M PBS solution. Due to the hybridization between the barcode DNA and its complements that are modified on Au-NPs surfaces, Au-NPs aggregate and show visible color change. And, the mixed solution, after being maintained at room temperature for 2 h, can be transferred into a 100-µL quartz cuvette to measure the extinction coefficients by using a UV-vis spectrophotometer [29,68].

Colorimetric Assay using Enzymes Labeled Au-NPs
Due to the high loading of signaling molecules on each nanoparticle, greatly enhanced sensitivity for the detection of tumor markers can be achieved by using enzyme labeled Au-NPs [70]. In such colorimetric assays, Au-NPs play a role of enzyme carriers and signal amplifiers, and the detectable signals come from the catalytic reaction of the enzymes that are loaded onto the nanoparticle surfaces. The commonly used enzymes and substrates are the same as those used in ELISA-based colorimetric assays.

Results and Discussion
As an example, one recent work in the authors' laboratory on colorimetric multiplexed immunoassays is introduced [45]. Figure 1 illustrates the principle of the assay for sequential detection of tumor markers. Briefly, biotin-anti-CEA conjugated MPs are first added to the test solution that contains CEA, AFP and some nonspecific proteins. After magnetic separation, CEA antigen which is captured by biotin-anti-CEA, immobilized on the surface of MPs, is thus collected. Therefore, if Au-NPs loaded with HRP-anti-CEA are added to the collected CEA antigen solution, HRP-anti-CEA loaded on the surface of Au-NPs may capture the CEA antigen conjugated on the surface of MPs. Consequently, when TMB and H 2 O 2 are added to the test solution, its color turns blue. Alternatively, the detection of AFP antigen in the test solution can be carried out by repeating the above procedure, using biotin-anti-AFP conjugated MPs and Au-NPs loaded with HRP-anti-AFP.
The multiplexed immunoassay is very simple and easily operated, and the results can be seen by the naked eye ( Figure 2). Meanwhile, the absorbance at 370 nm or 490 nm of the test solutions can be separately used to determine the amount of tumor markers more accurately. Under optical conditions, the immunoassay can provide sequential detection of CEA and AFP with detection limits of 0.02 ng/mL (≈0.1 pM), which can be satisfactory for clinical applications.  Colorimetric assay is very simple and easily operated, without requiring the expensive instruments needed in the optical immunoassay systems, such as charge-coupled device (CCD) camera and multi-channel injection valves fixed to luminescence analyzers. In fact, electrochemical assay is also very simple and relatively inexpensive, thus some electrochemical methods have also been proposed for the detection of cancer. However, the current amplification strategy for electrochemical signal often involves multiple steps of deposition and stripping, making the experimental process complex; while colorimetric approaches do not require complicated experimental steps, and results can be seen with the human naked eye. Therefore, colorimetric immunoassay for the detection of tumor markers has received more and more interest. Meanwhile, nanotechnology has greatly promoted the development of colorimetric immunoassays. Firstly, MPs may provide a rapid and safe approach to separate the target from the other species, preventing the interference of non-specific proteins in immunoassays. Secondly, the unique optical properties of Au-NPs can be harnessed to realize the transduction of the presence of target tumor markers to easily detectable signals, simplifying the designed detection system. Thirdly, the use of nanomaterials has enabled detection of tumor markers with greater sensitivity and accuracy. Therefore, colorimetric immunoassays have made prominent progress in the detection of tumor markers and may play an important role in the early diagnosis of cancer.
Colorimetric immunoassays have many desirable merits, thus are rapidly developing. Tremendous opportunities and challenges exist in the application of colorimetric methods for tumor marker detection.
Firstly, colorimetric immunoassays for tumor marker detection are usually proposed by using antibodies; however the availability of antibodies with high affinity and low cross-reactivity are very limited. Fortunately, aptamers-nucleic acid molecules that can selectively bind to low molecular weight organic or inorganic substrates or to macromolecules such as proteins-can be considered as attractive alternatives to antibodies [45]. In fact, compared with antibodies, aptamers may possess many advantages, such as relatively simple and inexpensive synthesis, tolerance to internal labeling, high stability, and stronger and more selective affinity for protein targets. Therefore, several aptamers have been employed for tumor marker detection [80,81].
Secondly, nanomaterials used in colorimetric methods for sensing tumor markers are still narrowly restricted to Au-NPs and MPs. Recently, some novel types of nanomaterials were fabricated and used in colorimetric biosensors for various analytes [82][83][84], so more nanomaterials will be applied in the development of colorimetric immunoassays for tumor marker detection, simplifying the assay methodology and lowering the detection limit as far as possible.
Finally, the measurement of a single tumor marker is usually not sufficient for diagnosis purposes, because most cancers have more than one marker associated with their incidence [85], thus multiplexed immunoassays that can quantitatively measure the concentrations of multiple tumor markers in a single assay have received more and more interest [25,46]. Compared with the traditional parallel single-analyte immunoassay, multiplexed immunoassays have the advantage of simplified analytical procedure, shortened analysis time, decreased sample consumption, increased test throughput, reduced costs, and improved diagnostic accuracy [45]. Consequently, future advances in colorimetric immunoassays will depend on the development of simple and efficient sensing mechanisms for multiplexed tumor marker examinations.