ABSTRACT
Leptospirosis caused by pathogenic Leptospira is one of the most common zoonoses in the world. It is believed that humans become infected with it mainly through their skin and mucous membranes by contact with water or soil that is contaminated with urine excreted from infected animals. Recently, outbreaks have frequently occurred in the tropics, especially after flooding, but how leptospires cause mass infection remains poorly understood. In this study, we injected leptospires into the tracheas of hamsters under direct view and prove for the first time that leptospires can infect through the respiratory tract. We determined that a 50% lethal dose (LD50) of the Leptospira interrogans strain UP-MMC-SM (L495) for hamsters in transtracheal infection was 3.2 × 102 cells. The results of culture, macroscopic findings, and histopathological analysis suggested that intratracheally injected leptospires invaded the lung tissue, proliferated in the collagen-rich stroma adjacent to the bronchus and blood vessels, and then spread throughout the body via the bloodstream. In the lung, leptospires continuously infiltrated the alveolar wall without inflammatory cell infiltration, spread throughout the lung, and finally caused pulmonary hemorrhage. Our results revealed that the respiratory tract might be a portal of entry for leptospires. We speculate that some cases of leptospirosis might be caused by transbronchial infection from inhaling infectious aerosols containing leptospires during floods. Leptospira was also confirmed to be a unique pathogen that invades through the bronchus, proliferates in the collagen-rich lung stroma, and spreads through the alveolar interstitium throughout the lung without causing pneumonia.
INTRODUCTION
Leptospirosis caused by Leptospira is one of the most common zoonotic diseases in the world, especially in tropical and subtropical areas with high temperatures and heavy rainfall (1, 2). There are two types of human leptospirosis: a mild, flu-like type without jaundice, and Weil’s disease, which is a severe type with jaundice, pulmonary hemorrhage, renal failure, and even death (1, 3, 4). Costa et al. (5) reported that leptospirosis was estimated to cause more than 1 million severe cases and about 60,000 deaths annually worldwide (5, 6). It has been believed that humans become infected mainly through their skin and mucous membranes by contact with water or soil contaminated with urine excreted from infected animals (7).
Leptospires are thin, helix-structured spirochetes with hook-like curved ends. Their width is as thin as 0.1 μm, and their length is 6 to 12 μm. One flagellum from each end passes through its periplasm between the inner membrane and the outer membrane and extends to the center of the bacterial cell. The flagellar structure maintains a characteristic helical structure. The entire bacterial cell rotates itself by rotating the flagella. Leptospires can move back and forth in a soft agar medium in a corkscrew-like manner. They are believed to be able to invade host tissues in the same manner (2).
The mechanism of the occurrence of outbreaks has not been fully elucidated, but in recent years, outbreaks of leptospirosis have occurred frequently, especially after flooding caused by heavy rain, hurricanes, and storm surges (3). Outbreaks also occur after sport and leisure activities in rivers and lakes (8–10). As mentioned above, infections through the mucocutaneous surface are considered to be the main route of infection in outbreaks, but it is unlikely that this is the route in all cases of outbreaks. During floods and sport and leisure activities in rivers and lakes, water contaminated with Leptospira may become aerosols and float in the air, and it is speculated that some cases of leptospirosis may be caused by infection through the respiratory tract by inhaling said infectious particles, as in legionellosis (11).
Hamsters, guinea pigs, and gerbils have been used in previous studies as experimental animal models of human leptospirosis because these animals are susceptible to leptospires and exhibit similar symptoms of Weil’s disease (jaundice, pulmonary hemorrhage, and renal failure) at the end stage of infection (12, 13). The infection routes used in the above-named animal models are usually either subcutaneous (14, 15) or intraperitoneal (13). To our knowledge, there have been no reports confirming whether or not leptospires can be transmitted through the respiratory tract in an animal model. That is probably due to the difficulty of the experimental technique for inoculating leptospires into the trachea.
In this study, we used the technique of injecting pathogenic leptospires into the tracheas of anesthetized hamsters under direct view to demonstrate respiratory tract infection as a novel route of Leptospira infection. We also performed a histopathological analysis of lung tissue infected with leptospires through the respiratory tract and compared it to an analysis of well-researched subcutaneous infection.
RESULTS
Survival of hamsters intratracheally infected with leptospires.Hamsters intratracheally infected with 2 × 100, 2 × 101, 2 × 102, 2 × 103, or 2 × 104 cells of Leptospira interrogans strain UP-MMC-SM (L495) (5 hamsters in each dose group) were monitored daily for changes in body weight and development of symptoms for 14 days. The hamsters did not show any symptoms, including respiratory symptoms, until 8 days postinfection. At 9 to 12 days postinfection, some infected hamsters showed weight loss, ruffled fur, and activity loss, became moribund, and were euthanized. The survival rates of the infected hamsters are shown in Fig. 1A. Intratracheal administration of 2 × 102 leptospires caused death in 40% of the hamsters, and 2 × 103 leptospires killed all of them. Leptospires were recovered from all the euthanized hamsters. The 50% lethal dose (LD50) of L. interrogans strain UP-MMC-SM (L495) by intratracheal administration was calculated to be 3.2 × 102 cells by the Behrens-Karber method. Thus, it was confirmed for the first time that leptospires can cause infection through the trachea and subsequent death in hamsters.
Survival (Kaplan-Meier curve) of hamsters with respiratory tract infection (A) and subcutaneous infection (B). (A) Survival of hamsters intratracheally infected with 2 × 100 cells, 2 × 101 cells, 2 × 102 cells, 2 × 103 cells, and 2 × 104 cells of strain UP-MMC-SM (L495) (5 hamsters in each dose group); (B) survival of hamsters subcutaneously infected with 1 × 100 cells, 1 × 101 cells, 1 × 102 cells, 1 × 103 cells, and 1 × 104 cells of strain L495 (5 hamsters in each dose group).
In order to compare results with those from subcutaneous infection, which has been used conventionally, 1 × 100, 1 × 101, 1 × 102, 1 × 103, or 1 × 104 cells of UP-MMC-SM (L495) were injected subcutaneously into the right inguinal region of hamsters (5 in each dose group). The survival rates of the subcutaneously infected hamsters are shown in Fig. 1B. As with hamsters with respiratory tract infection, some hamsters with subcutaneous infection showed weight loss, ruffled fur, and activity loss and became moribund at 9 to 12 days postinfection. The LD50 was 3.2 × 100 cells. Infected hamsters were observed to lose weight at 1 to 2 days prior to death, regardless of the infecting dose and infection route. The mean percentages of body weight loss prior to the death of the hamsters with respiratory tract infection and subcutaneous infection were 4.3% and 4.9%, respectively.
Systemic distribution of leptospires in the host after respiratory tract infection.We instilled a 200-μl leptospire suspension containing 2 × 104 cells of the UP-MMC-SM (L495) strain into the trachea of each hamster in order to analyze the systemic distribution of leptospires in the host after respiratory tract infection. Infected hamsters were euthanized on days 0 (i.e., 2 h) to 10 postinfection every day (3 per day). After observing the macroscopic findings in each organ, we calculated the numbers of leptospires in each homogenized organ (the right lung, right kidney, and a subsegment of the liver [segment 4]) and in fluid specimens (blood, urine, bronchoalveolar lavage fluid [BALF]) by the limiting dilution culture method on 96-well plates.
The changes in the numbers of leptospires in each organ and specimen after respiratory tract infection are shown in Fig. 2. In the early stage of infection (from 0 to 3 days postinfection), there were no macroscopic findings in the lungs, but leptospires were isolated from the lung and BALF (Fig. 2A and F). At 3 days postinfection, leptospires were also isolated from the liver of an infected hamster (Fig. 2B). In the middle stage of infection (from 4 to 6 days postinfection), no symptoms, such as weight loss, ruffled fur, dyspnea, or activity loss, were observed, but pulmonary hemorrhage was observed macroscopically at 6 days postinfection (Table 1). At this stage, leptospires were isolated from all the organs and specimens except the urine, and the number of leptospires increased day by day (Fig. 2A to D and F). The periods from intratracheal infection to the first isolation of leptospires increased in length in the following order: lung and BALF, liver, blood, and kidney. In the end stage of infection (at 7 to 9 days postinfection), weight loss, ruffled fur, and activity loss were observed. Pulmonary hemorrhage, renal hemorrhage, intestinal bleeding, and jaundice became macroscopically visible at 8 days postinfection (Table 1; Fig. 3). Leptospires were isolated from the urine at 8 days postinfection (Fig. 2E). Moribund hamsters appeared at 9 days postinfection.
Changes in the number of leptospires in each organ and specimen after respiratory tract infection by the limiting dilution culture method on 96-well plates. A 200-μl leptospire suspension containing 2 × 104 cells of strain UP-MMC-SM (L495) was instilled into the tracheas of 7-week-old male hamsters. Infected hamsters were euthanized on days 0 (i.e., 2 h) to 9 postinfection every day (3 per day). The number of leptospires in each homogenized organ (right lung [A], a subsegment of the liver [segment 4] [B], and right kidney [C]) and specimen (blood [D], urine [E], and BALF [F]) was calculated by the limiting dilution culture method on 96-well plates. The dots indicate the number of leptospires in each individual hamster. The dashed lines indicate the detection limit of leptospires.
Numbers of hamsters with macroscopic pathological findings on each day after intratracheal infection (3 per day)
Macroscopic lesions in hamsters intratracheally infected with strain UP-MMC-SM (L495). Pulmonary hemorrhage (A), renal hemorrhage (B), and intestinal hemorrhage and jaundice (C).
We concluded from the above that leptospiral proliferation in the lungs of the intratracheally infected hamsters occurred earlier than that in subcutaneously infected hamsters reported in a previous study (2). The infected animals survived pulmonary hemorrhage without showing any symptoms, but they showed activity loss immediately after renal hemorrhage occurred. It was not possible to determine whether the direct cause of death in the infected hamsters was renal failure or pulmonary hemorrhage.
Changes in the numbers and fractions of leucocytes in BALF.The numbers of leukocytes in the BALF were counted with a hemacytometer, and the fractions of leukocytes there were determined by Wright-Giemsa staining. The results are shown in Fig. 4. The numbers of leukocytes did not increase in the early or even in the middle stages of infection (0 to 6 days postinfection) despite the proliferation of leptospires in the lung. An increase in leukocytes and the appearance of neutrophils in the BALF were observed at the end stage of infection (7 and more days postinfection).
Changes in the numbers and fractions of leucocytes in the BALF of intratracheally infected hamsters. A 200-μl leptospire suspension containing 2 × 104 cells of strain UP-MMC-SM (L495) was instilled into the tracheas of 7-week-old male hamsters. Infected hamsters were euthanized on days 0 (i.e., 2 h) to 9 postinfection every day (3 per day). The number of leukocytes in the BALF was counted using a hemacytometer, and the leukocyte fractions in the BALF were determined by Wright-Giemsa staining. As a noninfected control, BALF was collected from hamsters intratracheally administered 200 μl of only PBS on day 0 (n = 2).
Leptospiral infiltration in the lung after respiratory tract infection and subcutaneous infection.In order to investigate the differences in levels of leptospiral infiltration in the lung tissue after respiratory tract infection and subcutaneous infection, hamsters infected by each infection route were euthanized at 0, 2, 4, 6, 7, 8, and 9 days postinfection (2 per day). The left upper lungs were collected from each sacrificed hamster, fixed with a mixture of paraformaldehyde and glutaraldehyde, cut into 4-μm slices (three serial sections), and then stained with hematoxylin and eosin (HE), Elastica van Gieson (EVG) stain, and immunofluorescence stain.
The results are shown in Fig. 5. No significant change was observed in the HE-stained lungs of the intratracheally infected hamsters from 0 to 4 days postinfection from those of the noninfected controls. Pulmonary hemorrhage was observed both microscopically and macroscopically from 6 to 9 days postinfection. The alveolar cavity was filled with blood cells, but no obvious destruction of the lung tissue (i.e., blood vessel breakage, etc.) was observed microscopically. In the immunofluorescence staining, only a few leptospires were found in the peribronchial tissue at 0 days postinfection. Accumulation of leptospires was observed in the stroma adjacent to the bronchus and blood vessels (pulmonary artery or vein) at 2 days postinfection. At 4 days postinfection, leptospires increased in number over time and continuously infiltrated the peripheral alveolar stroma (Fig. 5). Until the onset of pulmonary hemorrhage, no leptospires were observed in the alveolar space except on the first day of respiratory infection. The spiral structure characteristic of leptospires was confirmed by confocal laser scanning microscopy at 2 days postinfection (Fig. 6). EVG staining demonstrated that the accumulation sites of leptospires revealed by immunofluorescence staining were consistent with the collagen-rich sites stained with EVG staining (Fig. 5). Although the HE-stained slices were examined in detail again, no inflammatory cell infiltration or tissue destruction was observed in the accumulation sites of leptospires revealed by immunofluorescence staining (Fig. 5B).
Leptospiral infiltration in the lung after respiratory tract infection and subcutaneous infection. (A) Hamsters intratracheally infected with 2 × 106 cells of strain UP-MMC-SM (L495) were euthanized at 0, 2, 4, 6, 7, 8, and 9 days after infection (2 per day), and the left upper lungs were collected, fixed, cut into 4-μm slices (three serial sections), and then stained with HE, immunofluorescence stain, and EVG stain. Representative microscope images of the serial sections at 0, 2, 4, 6, and 7 days postinfection are shown. Hamsters subcutaneously infected with 1 × 106 cells of strain L495 were euthanized at 0, 2, 4, 6, 7, 8, and 9 days postinfection (2 per day), and the left upper lungs were collected, fixed, cut into 4-μm slices (three serial sections), and then stained with HE, immunofluorescence stain, and EVG stain. Representative microscope images of the serial sections at only 7 and 8 days postinfection are shown because no changes were observed on days 0 to 6 postinfection. For immunofluorescence staining, leptospires were stained with rabbit anti-Leptospira interrogans serovar Manilae antiserum and goat anti-rabbit IgG antibody labeled with Alexa Fluor 488 (green), and the nuclei of lung cells were stained with DAPI (blue). In EVG staining, elastic fibers and cell nuclei were stained black, collagen fibers were stained red, and other tissues were stained yellow. The areas boxed by black frames in the HE-stained images are enlarged in panel B. The area boxed by a white frame in the immunofluorescence stain of day 2 in panel B is enlarged in Fig. 6. Scale bars, 200 μm (A) and 20 μm (B).
Enlarged immunofluorescence stain image of lung tissue after respiratory tract infection. The area boxed by a white frame in the immunofluorescence stain of day 2 in Fig. 5B is enlarged. Leptospires were stained with rabbit anti-Leptospira interrogans serovar Manilae antiserum and goat anti-rabbit IgG antibody labeled with Alexa Fluor 488 (green), and the nuclei of lung cells were stained with DAPI (blue). The spiral structure characteristic of leptospires was confirmed. Immunofluorescence staining in the lung on day 2 postinfection was observed with a confocal laser scanning microscope. Scale bar, 20 μm.
The subcutaneously infected hamsters, on the other hand, showed the following symptoms and histopathological findings in the lungs. The infected hamsters exhibited body weight loss and macroscopic pulmonary hemorrhage at 8 days postinfection. There was no significant change from 0 to 7 days postinfection in the HE-stained lungs from those of the noninfected controls. It was not until 8 days postinfection that pulmonary hemorrhage could be observed microscopically, but there was no destruction of the lung tissue, even then. No leptospires were observed by immunofluorescence staining in the lung tissue at 0 to 7 days postinfection. Leptospires became diffusely located in the alveolar interstitium at 8 days postinfection (Fig. 5).
We concluded from the above that the stroma adjacent to the bronchus and the blood vessels were the first colonization and accumulation sites of leptospires in the lungs of the intratracheally infected hamsters. These sites were consistent with the collagen-rich sites stained by EVG staining. In the subcutaneously infected hamsters, on the other hand, there was no leptospiral colonization or proliferation in the lungs in the early and middle stages. Leptospires suddenly appeared in the peripheral alveolar interstitium at the end stage of infection. The degree of macroscopic pulmonary hemorrhage in the subcutaneous infection was milder than that in the respiratory tract infection (Fig. 5A).
Observation by TEM and SEM-CLEM of the location of leptospires at the end stage of infection.In order to observe in detail the locations of leptospires at the end stage of infection, the lower left lung collected from an intratracheally infected hamster at 7 days postinfection was analyzed by transmission electron microscopy (TEM). The lower right lung of the same infected hamster was cut by the cross-cutting method, and the surface was observed by scanning electron microscopy-correlative light and electron microscopy (SEM-CLEM).
As shown in Fig. 7, there were leptospires near the collagen fibers in the intercellular spaces of the peripheral alveolar stroma. Some leptospires were phagocytosed by alveolar macrophages, covered with a membrane, and swirling. In the SEM-CLEM photograph of Leptospira, the organism was observed on the surfaces of separated alveolar cells exposed by dissociation of intercellular adhesion by the cross-cutting method (Fig. 8), which was confirmed by fluorescence microscopy. The photograph in Fig. 8 indicates that leptospires penetrated through the gaps between the alveolar cells.
Transmission electron microscope (TEM) images of the lung at the end of intratracheal infection with leptospires. (A) The lower left lung at 7 days postinfection was analyzed by TEM. Leptospires (white arrow) were phagocytosed by alveolar macrophages, covered with a membrane, and swirling. (B) There were leptospires near the collagen fibers (white stars) in the intercellular spaces of the peripheral alveolar stroma. (B, C) Shadowing was observed around the leptospires, suggesting that the leptospire surface may have been coated with a substance. Scale bars, 5 μm (A), 1 μm (B), and 2 μm (C).
Scanning electron microscope-correlative light and electron microscope (SEM-CLEM) images of the lung at the end of intratracheal infection with leptospires. The lower right lung of an infected hamster at 7 days postinfection was cut by the cross-cutting method, and the surface was observed by SEM-CLEM. Leptospires were stained with rabbit anti-Leptospira interrogans serovar Manilae antiserum and goat anti-rabbit IgG antibody labeled with Alexa Fluor 488 (green) (images on the bottom left of each panel). The data taken with an SEM (images on the top left of each panel) after fluorescence staining were edited (images on the right of each panel) using Photoshop CC (overall view [A], enlarged view of the area surrounded by a white box in panel A [B]). Leptospira cells were observed on the surfaces of separated alveolar cells exposed by dissociation of intercellular adhesion with the cross-cutting method. Scale bars, 10 μm (A) and 1 μm (B).
DISCUSSION
In this study, using a hamster model, we confirmed for the first time that leptospires can infect through the respiratory tract. The 50% lethal dose (LD50) of L. interrogans strain UP-MMC-SM (L495) in transtracheal infection was determined to be 3.2 × 102 cells (Fig. 1A). Although this LD50 was about 100 times more than that of the same strain in subcutaneous infection (Fig. 1), it cannot be concluded that the leptospires infected more easily through the skin than through the trachea, because the leptospires in the experimental subcutaneous infection were injected by needle that had passed through the epidermal barrier.
Respiratory tract infection is considered to be an important route of leptospiral infection acquired from the environment (2, 8, 16, 17). Water contaminated with urine excreted from Leptospira-infected animals becomes an aerosol and floats in the air, and then humans and animals can be infected through their respiratory tracts by inhaling the infectious particles or by aspiration of contaminated water (8, 9).
Based on the results of the culture, macroscopic findings, and histopathological analysis in this study, we concluded that the leptospirosis caused by inhaled pathogenic Leptospira progresses as follows. First, the inhaled leptospires colonize the surfaces of bronchial mucosae and invade the collagen-rich stroma adjacent to the bronchus and blood vessels (pulmonary artery or vein), as shown by the immunofluorescence staining in Fig. 5. After invading the stroma, leptospires grow there to some extent and then spread throughout the body via the bloodstream. The colonized leptospires grow in each tissue and cause multiple-organ failure. As shown in Fig. 2, the periods from infection to the first isolation of leptospires increased in length in the following order: lung, liver, blood and kidney, and urine. This indicates that the time that leptospires need to move from the initial growth sites of the lungs to each organ and to grow there is in that order. In the lung, leptospires proliferate at the collagen-rich stroma adjacent to the bronchus and blood vessels and then continuously infiltrate the alveolar wall through the interstices in the stromata (Fig. 5). After leptospires begin to circulate in the bloodstream, some of them may return to the capillaries of the alveolar periphery from other organs, directly colonize the alveolar wall, and grow there.
The progress of systemic distribution of leptospires in subcutaneous infection, which has often been reported, was revealed to be different from that in respiratory tract infections. We previously investigated a hamster leptospirosis model by using an in vivo imaging system (IVIS) to observe the entire body of hamsters subcutaneously infected with Leptospira interrogans serovar Manilae strain M1307, which was the UP-MMC-SM (L495) strain with the luciferase gene. We showed that strain M1307 injected subcutaneously in the right inguinal region proliferated on the vascular endothelium of adipose tissue in the early stage of infection, after which the leptospires spread along the blood vessels of subcutaneous adipose tissue and were disseminated throughout the whole body via the bloodstream (15).
In this study, we showed that leptospires became detectable in the lungs of subcutaneously infected hamsters at 8 days postinfection (Fig. 5), appearing diffusely in the alveolar wall without accumulating in the alveolar peripheral stroma. Unlike in respiratory tract infections, there was no leptospiral accumulation and proliferation in the collagen-rich stroma adjacent to the bronchus and blood vessels in the subcutaneously infected hamsters in the early and middle stages of infection. The degree of macroscopic pulmonary hemorrhage in subcutaneous infection was milder than that in respiratory tract infection (Fig. 5A). We concluded that there are differences between respiratory tract infection and subcutaneous infection in the initial growth site of leptospires, the course of leptospiral infiltration into the alveolar wall, and the severity of pulmonary hemorrhage, as shown in Fig. 9.
Summary of the processes of leptospiral infiltration of the lungs of hamsters with respiratory tract infection and subcutaneous infection.
Pulmonary hemorrhage seems to occur by blood leakage from the capillaries in the alveolar wall into the alveolar space. Pulmonary hemorrhage is considered to occur by the direct effect of leptospires because it was observed after leptospires had infiltrated the alveolar wall stroma in the middle stages of infection, although there were no findings of endothelial cell injury or architectural damage in the capillaries of the alveolar wall (Fig. 5, 7, and 8). We previously showed by TEM that leptospires penetrated the intercellular adhesion of hepatocytes and caused jaundice by destruction of the bile canaliculi between the hepatocytes (14). Sato and Coburn (18) demonstrated that pathogenic Leptospira disturbed the biological structures of the adherens junctions (vascular endothelial cadherin [VE-cadherin] and catenin) to enhance vascular permeability. Based on these reports, we speculated that leptospires invade the endothelial cell gap to move into and out of capillary blood vessels, consequently break the adherens junctions, and cause pulmonary hemorrhage due to increased vascular permeability. Pulmonary hemorrhage was observed at 6 days postinfection, but no symptoms, such as weight loss, dyspnea, or activity loss, were observed. Hamsters became moribund immediately after renal hemorrhage was observed at 8 days postinfection. Although the infected hamsters did not die immediately after pulmonary hemorrhage occurred, there are previous reports of human patients with leptospirosis dying of hemorrhagic pneumonia (19–22). It was not possible, therefore, to determine whether the direct cause of death in our animal experiment was renal failure or pulmonary hemorrhage.
According to the textbook Pathologic Basis of Disease by Robbins and Cotran (23), the histologies of bacterial pneumonia vary with the type of pathogen. Common bacterial pneumonia caused by Streptococcus pneumoniae or Staphylococcus aureus, etc., shows purulent fibrin exudates. Atypical pneumonia caused by Mycoplasma pneumoniae, etc., shows interstitial infiltration of mononuclear cells. Chronic pneumonia shows granuloma species and hollowing. Bacterial pneumonia exhibits either a pattern of bronchopneumonia or lobar pneumonia. Bronchopneumonia spreads to the surrounding alveoli centered around the bronchus, is distributed in spots, and affects the lobes of the lung. Lobar pneumonia is uniformly filled with exudate from lung lobes or some alveoli. Histologically, both show capillary dilation in the acute phase and neutrophil infiltration with precipitation of fibrin in bronchial and alveolar spaces. Macrophage infiltration is observed with the progress of inflammation, and alveolar exudate is replaced by granulation tissue or fibrous tissue. Unlike with common bacterial pneumonia, the alveolar space is not filled with exudate in atypical pneumonia.
In the pathological findings of the present study, lung tissue infected with leptospires did not match any findings of common bacterial pneumonia, atypical pneumonia, or chronic pneumonia. HE staining showed no infiltration of inflammatory cells, despite the presence of numerous leptospires recognized by immunofluorescence staining in the lung tissue (Fig. 5B). In addition, as shown in Fig. 4, the number of leukocytes in the BALF did not increase in the early or even middle stage of infection, even though leptospires were present in the BALF. This suggested that leptospires may escape from the immune system in the early and middle stages of infection. An increase in leukocytosis and the appearance of neutrophils were observed in the BALF after the pulmonary hemorrhage occurred in the end stage of infection (Fig. 4). Leptospira organisms that flowed out into the alveoli in blood were phagocytosed by alveolar macrophages (Fig. 7A) (24). Matsui et al. (25, 26) showed that there was a delay in the expression of the proinflammatory cytokines interleukin 1β (IL-1β), IL-6, and tumor necrosis factor alpha (TNF-α) and the chemokines CXCL10/IP-1 and CCL3/MIP-1α and that a massive overexpression in Leptospira-infected hamsters contrasted with that strictly regulated in mice (asymptomatic carriers). The expression of the anti-inflammatory cytokine IL-10 was earlier and greater in mice than in hamsters (27). These findings indicated that in the early stage of Leptospira infection, the hamsters delayed production of cytokines, failed to activate lymphocytes and neutrophils, and allowed leptospires to escape from the host of the immune system.
There are two possible reasons why leptospires grow in the collagen-rich stroma in the early stage of respiratory tract infection. First, collagen itself might be a source of nutrients for leptospires to grow. Second, leptospires preferentially accumulate on the high-order structure of collagen. It has been reported that L. interrogans easily adheres to collagen (28) and that the production of the colA gene, encoding collagenase, is involved in invasion and propagation (29), but it has not been reported that leptospiral proliferation is promoted by collagen. To clarify whether or not collagen can be a source of nutrients for leptospires, we cultured L. interrogans strain UP-MMC-SM (L495) in Korthof’s medium mixed with type I collagen derived from bovine skin (acid soluble) (Nippi Co., Tokyo, Japan) and counted the number of leptospires over time. There was no significant difference in growth from that of the control (Korthof’s medium without type I collagen) (data not shown), leading us to suspect that leptospires accumulate to the high-order structure of collagen and proliferate there at the early stage of respiratory tract infection. Further studies are needed to determine the role of collagen in the initial growth of leptospires in the host.
In conclusion, this is the first study to prove that respiratory tract infection can be a route of leptospiral infection, by means of a hamster model. Our findings suggested that leptospires invaded the tissue through the intercellular space. The collagen-rich stroma of the lung was thought to be the site of the proliferation of leptospires in the early stage of respiratory tract infection, suggesting that collagen was involved in the initial proliferation. The leptospires in the lung might have escaped from the host’s immune system and did not cause inflammation or pneumonia in the early stage of infection, unlike many other pathogenic bacteria. It was also found that there were differences between respiratory tract infection and subcutaneous infection in the course of the spread of leptospires and in the levels of severity of pulmonary hemorrhage.
In order to prevent leptospiral infection, it is necessary not only to avoid contact with contaminated water but also to avoid inhalation of contaminated aerosols during flooding and in freshwater sports and leisure activities.
MATERIALS AND METHODS
Ethics statement.The animal experiments were reviewed and approved by the Ethics Committee on Animal Experiments at the University of Occupational and Environmental Health, Japan (permit number AE15-019). The experiments were carried out under the conditions indicated in the Regulations for Animal Experiments of the University of Occupational and Environmental Health, Japan, and law 105 and notification 6 of the Government of Japan.
Bacteria and animals.Leptospira interrogans serovar Manilae strain UP-MMC-SM (also known as strain L495), kindly provided by Yasutake Yanagihara, was maintained in animals at the National Institutes of Infectious Diseases by Nobuo Koizumi after it was isolated from the blood of a human patient with severe leptospirosis at the University of the Philippines in 1998 (15, 30, 31). The UP-MMC-SM (L495) strain was cultured at 30°C in Korthof’s medium without shaking (32). Bacterial cell counts were determined by dark-field microscopy using a Thoma counting chamber. Seven-week-old male golden Syrian hamsters (body weight, 100 g or more) (Japan SLC, Inc., Shizuoka, Japan) were used for the animal experiment.
Animal experimental infection.The hamsters with respiratory tract infection were anesthetized with sevoflurane and placed on a 60°-angled board by hanging them from a loop around their upper incisor teeth, as shown in Fig. S1 in the supplemental material (33). The larynxes of the hamsters were opened with an Optima Clx Miller blade 0 fiber for newborns (Timesco laryngoscope; Smith Medical, Japan) processed to fit the width of the mouths of the animals. A 21-gauge, 90-mm spinal needle (Unisis Co., Saitama, Japan) whose tip was rounded with a file was connected to a 1-ml syringe filled with a UP-MMC-SM (L495) culture diluted with phosphate-buffered saline (PBS) to a predetermined amount of bacteria. This spinal needle was inserted 4 to 5 cm into the tracheas of the hamsters under direct vision, and then 200 μl of leptospire suspension was injected into the trachea (Fig. S1). For comparison, 100 μl of UP-MMC-SM (L495) culture diluted with PBS to a predetermined amount of bacteria for each dose was injected subcutaneously into the right inguinal region of other hamsters with a 27-gauge needle. PBS alone was administered intratracheally or subcutaneously to other hamsters as a noninfected control for each experiment.
Determination of the lethal dose of leptospiral respiratory tract infection.Two hundred microliters of leptospire suspensions containing 2 × 100, 2 × 101, 2 × 102, 2 × 103, or 2 × 104 cells of low-passage-number (≤2 times with in vitro subcultures) strain UP-MMC-SM (L495) was instilled into the trachea of each hamster (5 in each dose group). In order to compare our results with those of conventionally used subcutaneous infection, 100 μl of a leptospire suspension containing 1 × 100, 1 × 101, 1 × 102, 1 × 103, or 1 × 104 cells of strain UP-MMC-SM (L495) was injected subcutaneously into the right inguinal region of other hamsters (5 in each dose group). The hamsters were monitored for 14 days after infection. As previously reported, hamsters that became moribund (presenting with symptoms such as weight loss, ruffled fur, and mobility loss) were anesthetized with sevoflurane and then euthanized by cervical dislocation. To confirm that leptospirosis was the cause of death, the organs of the sacrificed hamsters were homogenized and cultured using Korthof’s medium containing STAFF (sulfamethoxazole, trimethoprim, amphotericin B, fosfomycin, and 5-fluorouracil) (34). As a noninfection control, 100 or 200 μl of only PBS was administered subcutaneously or intratracheally to three hamsters. The 50% lethal dose (LD50) of strain UP-MMC-SM (L495) for hamsters was calculated by the Behrens-Karber method.
Analysis of the systemic distribution of leptospires in the host after respiratory tract infection.A 200-μl leptospire suspension containing 2 × 104 cells of strain UP-MMC-SM (L495) was instilled into the tracheas of hamsters by the method described above. Infected hamsters were euthanized 0 (i.e., 2 h) to 9 days postinfection every day (3 per day) for observation of the behavior of leptospires in the host after infection in the respiratory tract. Blood was collected from the orbital venous plexus of each hamster under anesthesia with sevoflurane. The rib cages of the anesthetized hamsters were opened and were perfused from the left ventricle with 30 to 50 ml of saline containing 0.5% heparin. Urine was collected from the bladder.
Bronchoalveolar lavage fluid (BALF) was collected by the following method. An indwelling venous needle (20-gauge, 32-mm Introcan Safety catheter; B. Braun Melsungen AG, Germany) was inserted into the trachea, banded with 3-0 thread (Alfresa Pharma Co., Osaka, Japan), and connected to a 10-ml syringe filled with 6 ml of saline. The saline was injected and aspirated repeatedly three times carefully so as not to damage the lung tissue, and the fluid collected last was used as BALF.
In order to count the number of viable leptospires in each organ and specimen, the right lung, right kidney, whole spleen, and a subsegment of liver (segment 4) were collected and homogenized in 2 ml of Korthof’s medium containing STAFF. Blood (20 μl), urine (20 μl), and BALF (20 μl) collected as mentioned above were transferred into 980 μl of Korthof’s medium containing STAFF. The number of leptospires in each homogenized organ and specimen was calculated by the limiting dilution culture method on 96-well plates. The plates were incubated at 30°C for 28 days, and the presence or absence of leptospires was observed with a dark-field microscope (14). The number of leukocytes in the BALF was measured with a hemacytometer (Sunlead Glass Co., Saitama, Japan), and the leukocyte fractions in the BALF were determined by Wright-Giemsa staining. As a noninfected control, BALF was collected from the hamsters that had been intratracheally administered 200 μl of PBS only (n = 2).
Analysis of leptospiral infiltration in the lung after respiratory tract infection and subcutaneous infection.A 200-μl leptospire suspension containing 2 × 106 cells of strain UP-MMC-SM (L495) was instilled into the tracheas of hamsters, as described above, and 100 μl of a leptospire suspension containing 1 × 104 cells of the same strain was injected subcutaneously into the right inguinal regions of another group of hamsters. These infected animals were euthanized on days 2, 4, 6, 7, 8, and 9 postinfection (2 per day). The rib cages of the infected animals were opened when the hamsters were under anesthesia with sevoflurane. They were perfused from the left ventricle with 30 to 50 ml of saline containing 0.5% heparin and subsequently fixed with a 30- to 50-ml mixture of 4% paraformaldehyde and 0.1% glutaraldehyde. The left upper lung was collected and fixed with 4% paraformaldehyde and 0.1% glutaraldehyde at 4°C for more than 7 days. The fixed left upper lung was paraffin embedded and cut into 4-μm slices with a microtome. The slices (three serial sections) were deparaffinized and stained with hematoxylin and eosin (HE), Elastica van Gieson (EVG) stain, and immunofluorescence stain, as described below.
Immunofluorescence staining.The slices of the fixed lung tissue were blocked with 3% bovine serum albumin in PBS (blocking buffer) at room temperature for 15 min and washed with PBS. The blocked slices were incubated overnight at 4°C with a rabbit anti-Leptospira interrogans serovar Manilae antiserum (1:200) as a primary antibody. After being washed with PBS, the slices were incubated at room temperature for 3 h with a goat anti-rabbit IgG antibody labeled with Alexa Fluor 488 (1:500; Life Technologies Co., Eugene, OR, USA) as a secondary antibody. After being washed with PBS, the slices were incubated at room temperature for 10 min with 2 μg/ml 4′,6-diamidono-2-phenylindole (DAPI) (1:1,000; Life Technologies Co., Eugene, OR, USA). After a final washing with PBS, a slide glass was mounted with SlowFade Gold antifade (Life Technologies Co., Eugene, OR, USA) and observed with a virtual slide system (VS120-L100-FL; Olympus Co., Tokyo, Japan) and a confocal laser scanning microscope (LSM880 with Airyscan; Zeiss Co., Germany) (14, 15).
TEM and SEM-CLEM.After being perfused and fixed as described above, the lower left lung was excised at 7 days postinfection, fixed with a mixture of 2% paraformaldehyde and 2% glutaraldehyde at 4°C for more than 7 days, and then cut into 1- by 1- by 1-mm pieces. The pieces were washed with PBS and fixed with 1% osmium tetroxide at 4°C for 1 h. After ethanol dehydration and acetone substitution, the temperature of the pieces was gradually raised from 40°C to 60°C with epoxy 812 resin (Nisshin EM Co., Ltd.), and polymerization was carried out for 3 days. The pieces were sliced to a thickness of 80 nm with a Leica EMUC7 ultramicrotome (Leica Microsystems Co., Germany) and stained with uranium acetate and lead stain. The slices were observed with a JEM-1200EX transmission electron microscope (TEM) (JEOL, Tokyo, Japan). The postinfection lower right lung was similarly excised on day 7, fixed with a mixture of 4% paraformaldehyde and 0.1% glutaraldehyde, and cut into 2- by 2- by 2-mm pieces by the cross-cutting method (14). The pieces were stained by immunofluorescence staining as described above and subsequently fixed with 4% glutaraldehyde at room temperature for 30 min. After acetone dehydration and tert-butyl alcohol substitution, the pieces were dried with a VFD-30 vacuum freeze dryer (Vacuum Device Ltd., Ibaraki, Japan) overnight. The dried pieces were fixed on an aluminum sample stand for scanning electron microscopy (SEM) and observed with a BX51 fluorescence microscope (Olympus Co., Tokyo, Japan). After being coated with osmium, the pieces were also observed with a JXA-8600MX SEM (JEOL, Tokyo, Japan). The data taken with the SEM and fluorescence microscope were edited using Photoshop CC (35, 36).
ACKNOWLEDGMENTS
We thank Ryoko Maekado and Katsuichi Onizuka for technical assistance. We also thank Christopher Carman for his valuable editorial advice on the manuscript.
This research received no specific grant from any funding agency in the public, commercial, or not-for-profit sectors.
FOOTNOTES
- Received 16 September 2019.
- Accepted 17 September 2019.
- Accepted manuscript posted online 23 September 2019.
Supplemental material for this article may be found at https://doi.org/10.1128/IAI.00727-19.
- Copyright © 2019 American Society for Microbiology.
