Comparisons of the topographic characteristics and ...

Journal of Microscopy, Vol. 00, Issue 0 2018, pp. 1–14

doi: 10.1111/jmi.12697

Received 27 November 2017; accepted 24 February 2018

Comparisons of the topographic characteristics and electrical charge distributions among Babesia-infected erythrocytes and extraerythrocytic merozoites using AFM , W . D E J . M E R C A D O - R O J A N O ∗, A . R U D O L P H ∗, J . W A N G ∗, J . M . L A U G H E R Y † & L. SCUDIERO∗ C . E . S U A R E Z †, ‡ ∗ Department of Chemistry and Materials Science and Engineering Program, Washington State University, Pullman, Washington, U.S.A.

†Program in Vector-Borne Diseases, Department of Veterinary Microbiology and Pathology, Washington State University, Pullman, Washington, U.S.A. ‡Animal Disease Research Unit, Agricultural Research Service, United States Department of Agriculture, Pullman, Washington, U.S.A.

Key words. Atomic force microscope (AFM), Babesia bigemina, Babesia caballi, Babesia bovis, frequency-modulated Kelvin probe force microscopy (FM-KPFM), merozoites.

Summary Tick-borne Babesia parasites are responsible for costly diseases worldwide. Improved control and prevention tools are urgently needed, but development of such tools is limited by numerous gaps in knowledge of the parasite–host relationships. We hereby used atomic force microscopy (AFM) and frequency-modulated Kelvin probe potential microscopy (FM-KPFM) techniques to compare size, texture, roughness and surface potential of normal and infected Babesia bovis, B. bigemina and B. caballi erythrocytes to better understand the physical properties of these parasites. In addition, AFM and FM-KPFM allowed a detailed view of extraerythrocytic merozoites revealing shape, topography and surface potential of paired and single parasites. B. bovis-infected erythrocytes display distinct surface texture and overall roughness compared to noninfected erythrocytes. Interestingly, B. caballiinfected erythrocytes do not display the surface ridges typical in B. bovis parasites. Observations of extraerythrocytic B. bovis, B. bigemina and B. caballi merozoites using AFM revealed differences in size and shape between these three parasites. Finally, similar to what was previously observed for Plasmodium-infected erythrocytes, FM-KPFM images reveal an unequal electric charge distribution, with higher surface potential above the erythrocyte regions that are likely associated with Babesia parasites than over its remainder regions. In addition, the surface potential of paired extraerythrocytic B. bovis Mo7 merozoites revealed an asymmetric potential distribution. These observations may be important to better

Correspondence to: L. Scudiero, Chemistry Department and Material Science & Engineering Program, Washington State University, Pullman WA 99164-4630, U.S.A. Tel: 509-335-2669; e-mail: [email protected]

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understand the unique cytoadhesive properties of B. bovisinfected erythrocytes, and to speculate on the role of differences in the distribution of surface charges in the biology of the parasites. Introduction Babesiosis is an emerging health threat of global impact that affects humans, domestic and wild animals (Suarez & Noh, 2011). Bovine babesiosis, mainly caused by B. bovis and B. bigemina, is widely recognised as a devastating tick-borne cattle disease occurring in tropical and semitropical areas worldwide, remaining as an important threat for the development of bovine industry globally (Suarez & Noh, 2011; Florin-Christensen et al., 2014). A similar disease occurring in horses is caused by the less studied B. caballi. Babesia parasites have a complex life cycle that includes intraerythrocyte stages in the vertebrate host and sexual stages in the midgut of their tick vectors. Sexual reproduction leads to the production of tick stages involved in transovarial transmission, a defining feature of Babesia parasites. Babesia bovis causes a severe clinical disease which is related to the invasion of the host erythrocytes and parasite sequestration in capillaries, triggering neurological manifestations similar to those observed in human patients with severe malaria, which is caused by the infection of virulent strains of Plasmodium falciparum (Allred & Al-Khedery, 2004; Suarez & Noh, 2011). Although the neurological disease occurring in cattle infected with B. bovis has been related to its ability to sequester in cerebral capillaries, the mechanisms involved remain poorly understood (Allred & Al-Khedery, 2004). However, the ability of B. bovis to sequester in capillaries has been associated to the expression of parasite proteins, such as members of the VESA gene family,

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concomitant with structural changes in the surface of the infected erythrocytes, including ridge-like structures (O’Connor et al., 1999, 2000; Allred et al., 2000; Gohil et al., 2010). In contrast, the intraerythrocytic B. bigemina does not cause neurological symptoms, capillary sequestration, and does not express ridges in the surface of infected erythrocytes (Gohil et al., 2010). Consistently, B. bigemina does not appear to express such VESA like antigens in the surface of infected erythrocytes. In addition, the ability of B. caballi parasites to sequester in horse capillaries has been described (Wise et al., 2013, 2014), but the mechanisms involved remain poorly characterised. Because sequestration mechanisms in malarial and in B. bovis parasites involve modifications on the surface of infected erythrocytes operated by the parasites, it would be important to determine whether such modifications also occur in B. caballi-infected cells, as a predictor of the ability for sequestration by iRBC. The structural changes caused by the parasite include the presence of ridge-like structures in B. bovis-infected erythrocytes, a type of surface modification that might provide increased traction or adhesion to the infected erythrocytes and may be analogous to the knobs identified on the surface of Plasmodium-infected erythrocytes (Gohil et al., 2010). The ridge-like membrane protrusions appear to facilitate the adhesion of B. bovis-infected RBCs to vascular endothelium cells. Therefore, cell surface characterisation is needed for a better understanding of the host–parasite interactions and ultimately may help guide the design of improved strategies for control. AFM has been previously used to study and compare Babesia-infected and -noninfected red blood cells (Aikawa et al., 1997; Nowakowski & Luckham, 2002; Hutchings et al., 2007; Gohil et al., 2010; de Souza & Rocha, 2011), but interactions between the parasites and the host, such as erythrocyte invasion and sequestration that leads to virulence remain poorly characterised in Babesia. One reason for these gaps in our knowledge is the paucity of practical and sensitive methods for analysing cell surfaces in native or nearly native state. AFM was demonstrated to be able to provide a direct insight on the topography of the surface of B. bovis, and B. bigemina-infected erythrocytes (Hutchings et al., 2007; Gohil et al., 2010), but the surface characteristics of B. caballi-infected erythrocytes remain unexplored. In addition, Kelvin probe force microscopy (KPFM) technique was first used to measure the contact potential between different materials (Nonnenmacher et al., 1991; Melitz et al., 2011), and also to measure the surface potential difference in the external membrane of Plasmodium-infected erythrocytes (Aikawa et al., 1996; Akaki et al., 2002). Differential surface membrane potential may also be related to important biological functions such as the ability of the parasite and infected erythrocytes to interact with other cells and their environments. This includes the ability of merozoites to recognise, attach and invade erythrocytes, allowing the invading merozoites to discriminate between infected and noninfected cells, and the ability of B. bovis-infected erythrocytes to attach and sequester to epithelial capillary cells.

Hereby, we describe the use of the AFM to image, study and compare the surface of B. caballi-infected erythrocytes with the previously characterised surfaces of erythrocytes infected with B. bovis (displaying ridges) and B. bigemina (not displaying ridges). We also obtained and compared AFM images of B. bovis, B. bigemina, and B. caballi extracellular merozoites for the first time. Finally, we analysed electrical charge distribution differences in the surface of B. bovis-infected erythrocytes and extraerythrocytic B. bovis merozoites using FM-KPFM. The findings could help reveal mechanisms operating in cell–cell interactions and may provide insights on the ability of the cells to invade and to sequester using cytoadherence mechanisms. Materials and methods Parasites and blood cell samples The B. bovis parasites were grown in long-term microaerophilous stationary-phase culture as previously described (Levy & Ristic, 1980; Hines et al., 1989) The T3Bo strain (B. bovis Texas S74-T3Bo derived from S1-T2Bo (Goff et al., 1998) and the Mo7 biological clonal strain of B. bovis (Rodriguez et al., 1983; Hines et al., 1989) were maintained as cryopreserved stabilates in liquid nitrogen (Palmer et al., 1982). The Puerto Rico B. bigemina and the B. caballi strains used in the studies were previously described (Hotzel et al., 1997; Schwint et al., 2008). The blood cell samples were prepared from a culture expansion in wells by diluting infected red blood cells 1:1 with HL-1 media for 3–4 days until parasitemia reaches a desired level (>20%). The blood was then smeared on glass slides by mixing 2 µL of media with 2 µL of infected red blood cells and the dry slides were then placed in a 90% ethanol solution for 24 h inside a 50 mL tube to wash off contaminating salts. Finally, the slides were air dry before use. Extraerythrocytic merozoite samples The extraerythrocytic merozoite samples were made from a culture expansion in wells by diluting infected red blood cells 1:1 with HL-1 media for 3–4 days until parasitemia reaches a desired level (>20%). The infected red blood cells were centrifuged at 400 × g in a microcentrifuge for 4 min to pellet RBCs. The pellets were then washed twice in 1× phosphate buffered saline (PBS) and centrifuge at 400 × g for 4 min. The red blood cell pellet was then lysed in 300 µL of Ammonium Chloride-Tris buffered solution. The samples were left to incubate for 5 min at room temperature (or until blood cells lyse). After lysis, 300 µL of 1× PBS was added and the samples centrifuged at 2800 × g for 4 min to pellet merozoites, washed with 1X PBS and centrifuge again at 2800 × g for 4 min to pellet merozoites. A 2 µL of material was smeared on a slide and left to dry in air. The dry slide was placed in methanol for 30 s without movement then dipped up and down for 90 s and air dried. C 2018 The Authors C 2018 Royal Microscopical Society, 00, 1–14 Journal of Microscopy

COMPARISONS OF THE TOPOGRAPHIC CHARACTERISTICS AND ELECTRICAL CHARGE DISTRIBUTIONS

Atomic force microscopy (AFM) and frequency-modulated Kelvin probe force microscopy (FM-KPFM) To image and quantify cells and free B. bovis merozoites a scanning probe atomic force microscope (DI) with a NanoScope IIIa controller (Veeco Metrology, Santa Barbara, CA, USA) was used. Probes used for ContactModeTM in air imaging were 100-mm-long, V-shape configuration gold-coated silicon nitride (Si3 N4 ) cantilevers with a tip radius of curvature less than 10 nm and spring constants ranging from 0.58 to 0.06 N m−1 . Both height and deflection images were captured at a resolution of 512 × 512 pixels and a scan rate of 1– 3 Hz depending on the scan scale, which ranged from tens of microns to 2 µm. The surface roughness values were obtained using an available commercial software ImageMetrology (SPIP, UK). The average roughness of an image is generally 1 M−1 N−1 defined as Sa = MN k=0 l = 0 |z(xk , yl )|, where the dimensional parameters M and N are equal for a quadrangular image, which is the case here. A Bruker NanoScope 8 was used to acquire the height and surface potential images. A frequency modulation KPFM (FMKPFM) detection technique was utilised because of its higher lateral resolution and accuracy to generate potential maps of the surface. A Pt tip (25Pt300B) was purchased from Rocky Mountain Nanotechnology (RMN). Typical settings for Potential are I and P gains of 2 and 5, respectively with a lift scan of 180 mV and drive amplitude of 3000 mV. A resolution of 512 × 512 and scan rate of about 1 Hz were used for all images. All the images were plane corrected using Global Leveling and filtered using Median 3 × 3 nondirectional noise, high and low values utilising a commercial scanning probe image processing, SPIP. Differences in cell size and roughness were assessed using a two-tail T-test. Nuclear stained cells and fluorescent microscopy B. bovis-infected RBC and extraerythrocytic merozoite samples obtained as previously described were nuclear stained by adding one drop of ProLongTM Gold antifade reagent with DAPI (Invitrogen) onto each sample smear and a cover slip was placed over the top. The samples were left to incubate for 5 min and visualised using Leica DMi8 inverted microscope with bright field and ultraviolet fluorescence. Images were processed using Leica LAS X analysis software. Results Comparison of features of Babesia bovis-, B. bigemina- and B. caballi-infected erythrocytes We first applied AFM imaging techniques to compare morphological and surface characteristics of infected erythrocytes in B. bovis, B. bigemina and B. caballi. Typical 3D AFM images of B. bovis-noninfected and -infected erythrocytes are shown in Figure 1(A). The control noninfected red blood cell C 2018 The Authors C 2018 Royal Microscopical Society, 00, 1–14 Journal of Microscopy

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present in B. bovis cultures has an average diameter of 5.31 ± 0.37 µm (Table 1). The normal erythrocyte size for bovine species varies between 5 and 6 µm (Adili et al., 2014), consistent with the size calculated by AFM in this study. The surface texture appears smooth with an average roughness (Sa) of 0.74 nm ± 0.07 nm, as determined on height images by using the commercial software SPIP on 0.5 × 0.5 µm2 area at the centre of the cell (Table 1). The size of erythrocytes infected with Mo7 and TX-virulent B. bovis shows diameters of 5.95 ± 0.37 µm and 5.94 ± 0.33 µm, respectively (Table 1), which corresponds to an increase of about 12 % for both the Mo7 and the T3Bo virulent-infected erythrocytes. In addition, the B. bovis-infected erythrocytes exhibit several previously described prominent features (Nowakowski & Luckham, 2002). Consistently, previously published AFM images featuring erythrocytes infected with distinct Babesia parasites include depressions (Hutchings et al., 2007; Gohil et al., 2010; de Souza & Rocha, 2011), which are not evident in AFM images of noninfected erythrocytes (Fig. 2). The average dimensions in length and depth of the depressions as measured by AFM for both Mo7 and T3Bo B. bovis-infected cells (1.70 ± 0.08 and 1.73 ± 0.15 µm in length; 0.43 ± 0.06 and 0.39 ± 0.04 in depth respectively) are shown in Table 1. We calculated the Sa estimated in an area of 0.5 × 0.5 µm2 of the ‘flat’ region of the surface, away from the depressions in five B. bovis-infected erythrocytes. Similar calculations were also performed using five noninfected erythrocytes. The Sa values measured are 1.54 ± 0.33 nm for the Mo7 and 1.13 ± 0.05 nm for TX virulent B. bovis parasites (Table 1). The calculated roughness of the surface around the depression doubles for Mo7 B. bovis parasites and increases only by 50% for the TX virulent cells when compared to 0.74 ± 0.07 nm for the noninfected erythrocytes. The difference in texture and Sa values between these erythrocytes can be mostly attributed to the presence of previously described small and randomly oriented ridges on the surface of the erythrocytes parasitised by B. bovis (see below) (Hutchings et al., 2007; Gohil et al., 2010). In Figure 1(B) we show 3D AFM images of noninfected and infected erythrocytes for B. bigemina collected from the same in vitro culture source. The average diameter of the cells are 5.43 ± 0.22 µm and 6.35 ± 0.46 µm for noninfected bovine erythrocytes present in the B. bigemina cultures and for B. bigemina-infected erythrocytes, respectively. The calculated Sa of the infected erythhrocytes increases by more than 47% compared to the roughness of the noninfected erythrocytes with values of 0.78 ± 0.08 nm and 1.15 ± 0.09 nm for normal and infected erythrocytes, respectively. However, and in agreement with previous observations (Hutchings et al., 2007), ridge-like or knob structures are not at all evident in the surface of B. bigemina-infected erythrocytes (Fig. 2D). Similar to B. bovis pronounced depressions are also seen in the infected cells. The dimension of the depressions is about 1.88 µm in length and 0.54 µm deep. In Figure 1(C) we show 3D AFM images of both normal and infected B. caballi

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Fig. 1. Three-dimensional (3D) AFM height images of control and infected erythrocytes. (A) noninfected bovine erythrocytes, Mo7 and T3Bo strains of B. bovis; (B) noninfected bovine and B. bigemina-infected erythrocytes and (C) noninfected equine and Babesia caballi-infected erythrocytes. Table 1. Averages and standard deviations for diameters for 8–10 isolated cells and 0.5 × 0.5 µm2 surface roughness (Sa), lenght (L) and depth (D) of the depressions for B. caballi, B. bigemina and B. bovis cells (diameters were obtained by using SPIP Particle and Pore Analysis feature). B. caballi

Diam. (µm) (No of cells) Sa (nm) (N° of cells) L (µm) D (µm)

B. bigemina

B. bovis

Normal erythocytes

Infected erythocytes

Normal erythocytes

Infected erythocytes

Normal erythocytes

M07 erythocytes

T3Bo virulent erythocytes

4.82 ± 0.36 (8) 0.75 ± 0.15 (5)

6.56 ± 0.38* (8) 1.38 ± 0.21* (5) 2.33 ± 0.49 0.31 ± 0.03

5.43 ± 0.22 (8) 0.78 ± 0.08 (5)

6.35 ± 0.46* (8) 1.15 ± 0.09* (5) 1.88 ± 0.25 0.54 ± 0.05

5.31 ± 0.37 (10) 0.74 ± 0.07 (5)

5.95 ± 0.37* (10) 1.54 ± 0.33* (5) 1.70 ± 0.08 0.43 ± 0.06

5.94 ± 0.33* (8) 1.13 ± 0.05 * (5) 1.73 ± 0.15 0.39 ± 0.04

* Significantly different compared to normal erythrocytes (p < 0.01).

erythrocytes. The normal size of the horse erythrocytes is considered to be 5.7 µm, which is larger than the size calculated in this study (4.82 ± 0.36 µm). Similar to what was described for B. bovis and B. bigemina, the B. caballi-infected equine erythrocytes also display depressions and an increase in diameter of about 30% compared to the control cells, but, and consistent with previous observations (Hutchings et al., 2007), neither

ridges nor knob-like structures are evident in the surfaces of the infected erythrocytes on inspection using AFM (Figs. 2E and F). The dimensions of the depressions are about 2.33 µm in length and 0.31 µm deep at the darkest region of the depression. A Sa value of about 1.38 nm ± 0.21 nm was measured for the infected cells which shows an increase of about 86% compared to noninfected horse erythrocytes (Table 1). The C 2018 The Authors C 2018 Royal Microscopical Society, 00, 1–14 Journal of Microscopy


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Fig. 2. AFM deflection images displaying the surface textures of (A) noninfected bovine erythrocytes, (B) B. bovis Mo7, (C) B. bovis T3Bo and (D) B. bigemina-infected bovine erythrocytes. Panel (E) depicts a noninfected equine erythrocyte and (F) a B. caballi-infected erythrocyte.

average measured roughness of noninfected equine eryhrocytes appears to be similar to the bovine erythrocytes with an average value of 0.75 nm ± 0.15 nm (Table 1). Similar to the B. bovis- and B. bigemina-infected erythrocytes (Table 1), the B. caballi-infected cells also have increased calculated roughness. Diameter, roughness, length and depth of depressions values for all cells, noninfected and infected B. bovis, B. bigemina and B. caballi, are shown in Table 1. C 2018 The Authors C 2018 Royal Microscopical Society, 00, 1–14 Journal of Microscopy

Detailed imaging of the surface of infected erythrocytes To better compare the surface texture among noninfected bovine and equine erythrocytes with those infected with B. bovis, Mo7 and TX virulent, B. bigemina and B. caballi, we generated high-resolution surface deflection images of the erytrocytes using AFM. The images are zoomed in to display a region of the infected cells including one depression

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Fig. 3. (A) 2D and 3D AFM height images of extraerythrocytic B. bovis merozoites. (i) Paired and (ii) single merozoites. 3D zoomed image of a pair of merozoites (iii). A line profile at the centre and along the merozoite in (ii) exhibits topographic details of the surface. (B) Confocal microscopy images of (i) intact B. bovis T3Bo-infected erythrocytes and extracellular merozoites and (ii) T3B extracellular merozoites obtained after erythrocyte lysis. Bright field, DAPI-stained and merged images are shown.

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Fig. 4. AFM height image of an extraerythrocytic Babesia bovis merozoite.

(Figs. 2A–F). Randomly distributed ridges, characteristic of the surface of B. bovis, are evident on the surface of the Mo7 and T3Bo B. bovis-infected erythrocytes (Figs. 2B and C). The AFM images showing patterns of erythrocyte surface modification caused by these parasites are essentially identical to previously described AFM images of B. bovis-infected erythrocytes (Hutchings et al., 2007; Gohil et al., 2010). The length of the ridges range from 120 to 200 nm in erythrocytes infected by the Mo7 and the T3Bo virulent strains (Figs. 2B and C, respectively). Consistent with previous observations (Hutchings et al., 2007; Gohil et al., 2010) such ridges, or other similar erythrocite surface structures, were not found to occur in noninfected erythrocytes (Fig. 2A) nor in B. bigemina-infected erythrocytes (Fig. 2D). Likewise, neither the surface of noninfected horse erythrocytes (Fig. 2E) nor horse erythrocytes infected with B. caballi (Fig. 2F) display any pronounced detectable ridge-like or Plasmodium-like knob structures (Hutchings et al., 2007; Gohil et al., 2010).

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the extracellular structures shown in Figure 3(B) confirms the nature of these structures as extracellular merozoites. 2D and 3D AFM images of paired merozoites display a boat shape with distinctive features on the surface (Fig. 3A, ii). AFM inspection of identical preparations performed on noninfected erythrocytes did not result in the identification of such boatshaped structures. Bright red colour in the picture displayed in Figure 3(A), Panel (ii) reveals high points (crests), and orange valley regions existing in between the crests and at the centre of the parasite as shown by the line profile through the parasite (Fig. 3A, iii). The revealed topography exhibits a broad high region at the wider part of the parasite, a depression followed by steps height increase to reach a narrow high height toward the apical pole of the parasite (Fig. 3A, ii and iii). The dimensions of B. bovis extracellular merozoites as measured by AFM are about 2.8 µm long and 90 nm high. The width is 0.85 µm at the wider region and 0.46 µm at the narrow region of the parasite. The shape of the merozoites is a feature that likely influences its fitness and ability to glide (Asada et al., 2012) and rapidly invade erythrocytes. Detailed topographic features of an extraerythrocytic B. bovis merozoite are also shown in Figure 4. Several high regions (bright colour) and low regions in between (darker colour) are displayed in this AFM image. To the best of our knowledge, this is the first report of an image of merozoites using AFM displaying its intricate tridimensional cell structure. A couple of typical B. bigemina extraerythrocytic merozoites are also shown in Figure 5(A). The B. bigemina and B. bovis merozoites have a similar shape, with comparable length but slightly smaller height (80 nm). A line profile drawn at the centre of one of the merozoite and through the long side of the merozoite revealing details of the topography of the surface is shown in Figures 5(B) and (C). Extracellular B. caballi merozoites are displayed in Figure 6(A). Upon more detailed examination (Fig. 6B), a typical B. caballi merozoites exhibit a more rounded shape with average length of 3.1 µm along the longer side of the merozoites. This parasite exhibits a more even surface with less dramatic topography (Fig. 6C) compared to B. bovis and B. bigemina. The average height of extracellular B. caballi merozoites as measured by AFM is about 90 nm.

Imaging extracellular Babesia parasites In addition to imaging the surfaces of infected red blood cells, AFM-based methods allow the close up imaging of unfixed extracellular merozoites for the three distinct Babesia parasites analysed in this study. The AFM image in Figure 3(A) shows extraerythrocyte structures morphologically resembling B. bovis free merozoites that were obtained after lysis of Mo7-infected erythrocytes. Extracellular merozoites can also be observed before (Fig. 3B, Panel i) and after lysis (Fig. 3B, Panel ii) in confocal microscopy images. In addition to their typical morphology, the confocal microscopy images generated using DAPI nuclear staining of C 2018 The Authors C 2018 Royal Microscopical Society, 00, 1–14 Journal of Microscopy

Kelvin probe force microscopy studies Frequency modulation Kelvin probe force microscopy (FMKPFM) enables imaging of the surface potential on materials by measuring a contact potential difference between the sample surface and the AFM tip. This technique was first used by Aikawa to investigate the membrane knobs of unfixed Plasmodium falciparum-infected erythrocytes (Aikawa et al., 1996). Other studies of surface potential involved indicators such as negatively charged and positively charged particles to probe the surface potential of samples. For instance, Akaki investigated the surface charge of plasmodium falciparum

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Fig. 5. 2D and 3D AFM height images of extraerythrocytic B. bigemina merozoites. Paired and single merozoites are seen in (A). A 3D zoomed image of a pair of merozoites (B). A line profile at the centre and along the merozoite exhibits topographic details of the surface (C).

merozoites by surface potential spectroscopy (SPS) and transmission electron microscopy (TEM) using indium tin oxide glass slide (ITO) as conducting base substrate and negatively charged nanogold and positively cationised ferritin particles to probe erythrocytes and merozoites surfaces (Akaki et al., 2002). Akaki’s work demonstrated that the negatively charged particles attached to the apical end of merozoites and positively charged particles to the entire surface of the erythrocytes. Danon’s work on treated rabbit red cells using cationised ferritin showed that the number of particles attached to the red cells was in good agreement with the results obtained by electrophoresis (Danon et al., 1972). Sherman & Greenan Jane (1986) used a similar labelling to investigate anionic residues distribution on the surface infected cells with plasmodium falciparum using TEM. Here, a nonconducting typical glass slide, no charge labelling particles were used to measure the surface potential difference. A platinum tip is first scanned over noninfected erythrocytes, and then over similarly prepared Babesia-infected erythrocytes to determine the surface potential difference between the depressions found in parasitised erythrocytes and the remainder of the cell. The FM-KPFM images reveal that the depressions have a more positive charge distribution com-

pared to the remainder of the red cell plasma membrane. Figure 7 displays both a height image (Figs. 7A and C) and its associated surface potential image (Figs. 7B and D) of T3Bo B. bovis parasites. The incomplete bright circle observed around the cells is most likely a scanning artefact as a line profile across the cell display an asymmetric overshooting feature with intensity dependent on the position of the line. At the location of the depressions symmetrical brighter areas are seen with an increased surface potential of about 200 mV above the remainder of the cell. Those brighter areas located in the same regions in both the height and surface potential images confirm that the more positive potential is due to a sudden change in morphology produced by the presence of the parasites. Surface potentials were also measured between the depression regions and the remainder of the cell in B. bigemina-infected erythrocytes (Fig. 8) and in B. caballi-infected erythrocytes (Fig. 9). Similar to what was found for bovine erythrocytes infected with B. bovis, the areas of the erythrocyte surfaces associated with the depressions show a more positive surface potential than the remainder of the cells and the noninfected red blood cells (approximately 120–160 mV). Thus, the patterns of surface potentials measured for B. bigemina-, B. bovisand B. caballi-infected erythrocytes are relatively similar. C 2018 The Authors C 2018 Royal Microscopical Society, 00, 1–14 Journal of Microscopy


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Fig. 6. 2D and 3D AFM height images of extraerythrocytic B. caballi merozoites. Paired and single merozoites are clearly seen in (A). A 3D zoomed image of a pair of merozoites (B). A line profile at the centre and along the merozoite exhibits topographic details of the surface (C).

The surface potential of extracellular B. bovis Mo7 merozoites were also measured (Fig. 10B) and compared with a height image of the same merozoites (Figs. 10A and C). Interestingly, the height image shows decreasing height towards the apical end of the cell, which matches a relatively more positive potential for the same area (about 100 mV) as seen in the potential profile in Figures 10(C) and (D). It is noteworthy to point out that the support used for these measurements is a typical nonconductive glass slide, which does not allow direct comparison with the results obtained by Akaki et al. Nonetheless, the results shown here also demonstrate an asymmetric charge distribution between the bulk of the merozoite and its apical end. Collectively, these patterns of asymmetric charge distribution may suggest electrical interaction as a possible mechanism operating between merozoites and red blood cells. Discussion The occurrence of dramatic alterations in the surface of infected erythrocytes was previously described to occur for Plasmodium and Babesia parasites. Whereas P. falciparuminfected erythrocytes display knobs, infected P. gallinaceum erythrocytes display furrows, but only P. falciparum-infected C 2018 The Authors C 2018 Royal Microscopical Society, 00, 1–14 Journal of Microscopy

erythrocytes have a cytoadhesive phenotype (Nagao & Dvorak, 1998). Ridge-like structures similar in morphology to the furrows found on the surface of infected P. gallinaceum erythrocytes were also found present in cytoadhesive B. bovis-infected erythrocytes. Consistently, among the three different Babesia parasites analysed in this study, B. bovis-infected erythrocytes uniquely exhibit ridge-like features that are pronounced and randomly distributed around the infected region of the cells. Both features, changes in size and surface characteristics, may have an impact in the host–parasite relationships. In addition, previous observations demonstrated that Babesia-infected erythrocytes are more rigid than non-infected erythrocytes (Hutchings et al., 2007). Nonetheless, of the cytoadherence mechanisms previously described for B. bovis, it is possible that these features may enhance sequestration or prolong the time of residence of Babesia-parasitised erythrocytes in capillaries, because it would be more difficult for erythrocytes to squeeze through narrow capillaries that may be as small as approximately 3 µm wide. Increased retention of parasitised erythrocytes in capillaries might help the parasite to avoid fast clearance by the spleen, which can contribute to pathogenesis and persistence of the infection. The calculated sizes for the length of the ridges in B. bovis-parasitised erythrocytes

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Fig. 7. FM-KPFM images of noninfected (Panels A and B) and B. bovis-infected (Mo7 strain) (Panels C and D) erythrocytes. Panels A and C: height images and Panels B and D surface potential images.

reported in this study for erythrocytes infected with the Mo7 strain is consistent with previous reports by Hutchings (Hutchings et al., 2007), who described the size of ridges ranging in between 140 and 250 nm for Australian strains of B. bovis. The lack of protruding ridge or knob like structures in B. bigemina is also consistent with the previous report by Hutchings (Hutchings et al., 2007). This difference in erythrocyte surface texture is also expected to have an important effect on the cytoadherence to plasma membrane of capillary vessels. As previously suggested, increased traction and expression of parasite proteins in the ridges may play a role in the increased cytoadherence or sequestration of B. bovis-infected erythrocytes to epithelial cells residing in the capillary vessels (Aikawa et al., 1996; Hutchings et al., 2007; Gohil et al., 2010). Parasitised erythrocyte sequestration, especially in capillaries of the brain and kidney, is the watermark of acute B. bovis infection in cattle. Furthermore, sequestration in brain capillaries leading to inflammatory responses likely contributes to the occurrence of neurological signs. Similarly, kidney failure in B. bovis-infected animals can also be related to sequestration of parasitised erythrocytes in the kidney’

capillaries. Interestingly, our data indicates that erythrocytes infected with the biologically cloned and poorly virulent line Mo7 contain higher surface roughness than the erythrocytes infected with the T3Bo virulent strain, thus at least in this case, virulence cannot be directly associated with the number of ridges present in the infected erythrocyte surface among these two parasites, and the data is in apparent conflict with previous observations correlating increased virulence with increased erythrocyte roughness (Hutchings et al., 2007). However, the Mo7 strain, which is considered as having an attenuated virulence phenotype, was not generated by the typical attenuation procedure involving serial passage of a virulent strain through a series of splenectomised steers (Suarez et al., 2012). Instead, the Mo7 clonal line was derived from a virulent strain using biological cloning by dilution in in vitro cultures (Rodriguez et al., 1983), and the differences in virulence may be accounted by differences in the pattern of expression of other and still unknown virulence factors. In addition, and consistent with previous findings, the ridged surface B. bovis-infected erythrocytes is distinct from the surface of the non-cytoadherent B. bigemina-infected C 2018 The Authors C 2018 Royal Microscopical Society, 00, 1–14 Journal of Microscopy


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Fig. 8. FM-KPFM images of noninfected (Panels A and B) and B. bigemina-infected (Panels C and D) erythrocytes. Panels A and C: height images and Panels B and D: surface potential images.

Fig. 9. FM-KPFM images of (A) height and (B) surface potential of infected B. caballi-infected erythrocytes.

erythrocytes (Gohil et al., 2010). In this study we show that this feature is also shared with the ridge-less B. caballi-infected erythrocytes. Intriguingly, the apparently featureless surfaces of B. bigemina- and B. caballi-infected erythrocytes also have increased values of their calculated Sa roughness (Table 1) compared to equally treated noninfected bovine and equine C 2018 The Authors C 2018 Royal Microscopical Society, 00, 1–14 Journal of Microscopy

erythrocytes, a feature that requires further investigation. It has been shown, for instance, that B. bigemina-infected erythrocytes uniquely contain host IgM molecules attached on their surfaces (Echaide et al., 1998). This feature can also modify the cell surface and provide increased roughness, as well as distinct morphological and physicochemical changes,

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Fig. 10. Height and potential FM-KPFM images of extraerythrocytic B. bovis Mo7 merozoites (Panels A and B). Panel C displays an asymmetrical height profile and the potential profile shown in Panel D reveals a larger potential at the centre of the parasite than at the apical pole.

which remain uncharacterised and may also impact the development of persistent infection by this parasite. The differences in infected erythrocyte architecture might also impact the clearance of infected parasites occurring mainly by spleen resident macrophages. It is possible that increased modifications in the surface of B. bovis-infected erythrocytes result in distinct rates of macrophage recognition of parasitised erythrocytes affecting the rate of parasitised erythrocyte clearance by the spleen. As in the case of B. bigemina-infected erythrocytes, increased surface roughness can not be correlated with the presence of ridges or other visible protruding structures in the surface of B. caballi-infected equine erythrocytes, and the nature of the erythrocyte surface modifications in B. caballi-infected cells responsible for increased Sa compared to noninfected erythrocytes needs to be further investigated. Consistently, all AFM images from erythrocytes infected with Babesia parasites in this study include large depressions, which are not evident in noninfected erythrocytes which can not be detected using other methods of imaging. In fact, our AFM images of B. bovis- and B. bigemina-infected erythrocytes are essentialy identical to what was previously reported, and

similar depressions were previously found in Plasmodiumand Babesia-infected erythrocytes in other AFM studies using a different sample preparation conditions (Hutchings et al., 2007; Gohil et al., 2010; de Souza & Rocha, 2011). Infected erythrocytes present either single or a double depression, depending on the parasite stage and the number of parasites resident in the infected cells. The trophozoite is a single cell stage occurring in Babesia-infected erythrocytes, and thus parasites with a single depression likely harbour trophozoite stage parasites. These depressions, if not artefactual but indeed caused by the parasite, are an intriguing erythrocyte surface modification. However, it remains unknown whether such depressions actually occur in in vivo blood-circulating infected erythrocytes, and the biological significance of this surface feature, if any, remains unknown. For the first time, AFM images of merozoites have been resolved to show common features and differences between Babesia parasites. The B. caballi merozoites are rounded whereas B. bovis and B. bigemina are boat shaped. Extracellular merozoites are known to glide following egression from infected erythrocytes until they find and invade a new C 2018 The Authors C 2018 Royal Microscopical Society, 00, 1–14 Journal of Microscopy


erythrocyte target (Asada et al., 2012). It is likely that these steps need to occur quickly enough to avoid recognition and binding of the immune response effectors of the host, such as antibodies and complement. The images suggest that extracellular merozoites have a complex surface and shape which likely evolved to optimise gliding and the mechanisms involved in erythrocyte recognition and invasion, as well as mechanisms for evading and minimising the effect of binding antibodies directed against merozoites surface molecules, which are typically found in persistently infected animals (Suarez & Noh, 2011). Finally, we hereby report the use of FM-KPFM techniques for the first time on Babesia-infected erythrocytes and merozoites for the measurement of the surface potential of both erythrocytes and merozoites. Interestingly, the infected regions of the erythrocytes occupied by the Babesia parasites exhibit a more positive potential than the remaining of the cell. These results are in good agreement with studies in which positively and negatively charged labelling particles and conducting substrate were used to measure the surface potential of the blood cells (Danon et al., 1972; Akaki et al., 2002). Free merozoites also display a similar surface potential, whereas the erythrocyte target cells are known to possess an oppositely charged surface, likely due to the expression of negatively charged sialylated glycoproteins (Fernandes et al., 2011), nanogold particles (Akaki et al., 2002). The difference in potential found on the surface of merozoites is large enough to bring merozoite and the target erythrocyte cells together before chemical binding takes place. It is thus possible to speculate that the initial interactions among free merozoites and noninfected erythrocytes may involve electrical interactions. Conversely, these measurements suggest that it is somehow possible that a similar electrically mediated repulsion mechanism would prevent or discourage free merozoites to interact with erythrocytes, which are already infected by the parasites. Further studies are needed to fully understand the significance of the difference in potential between the parasites and the remaining of the cells for the infected erythrocytes, including the possible role of surface exposed, highly glycosylated, IgM molecules, or any other modification caused by the parasite in the surface of the erythrocyte (Echaide et al., 1998). A possible interaction mechanism occurring between erythrocyte and merozoite, leading to the invasion of the erythrocyte by the parasite, could be the electrical attraction generated by the potential difference between cell and parasite before chemical binding happens. In summary, combined use of AFM and FM-KPFM provided experimental evidence of differential morphological and physicochemical characteristic of the surface of Babesiainfected erythrocytes and extracellular merozoites. The data discussed in this study may contribute to our better understanding of the interactions at play between erythrocytes and merozoites, the parasite host interactions, and it can be used to formulate and test novel experimental hypotheses. C 2018 The Authors C 2018 Royal Microscopical Society, 00, 1–14 Journal of Microscopy

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Conclusion (1) The surface of B. caballi-infected erythrocytes does not present the ridges typical in B. bovis-infected erythrocytes, despite significant roughness compared to noninfected erythrocytes. (2) Unequal electric charge distribution was identified in the surfaces of B. bovis-infected erythrocytes and extracellular merozoites. (3) Asymmetrical patterns of surface potential distribution was identified among B. bovis-infected erythrocytes and extracellular merozoites, suggesting the possibility of electrical charge-dependent mechanisms involved in interactions among cells. Acknowledgements We are grateful to Paul Lacy for his contribution in the production and preparation of the parasites used in the studies. We also thank Dan Riscioli, David A. Schneider for fruitful discussions and Lowell Kappmeyer for the provision of B. caballi parasites. The work described hereby was supported by the United States Department of Agriculture-Agriculture Research Service Current Research Information System Project No. 5348-32000-028-00D.

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C 2018 The Authors C 2018 Royal Microscopical Society, 00, 1–14 Journal of Microscopy