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. Author manuscript; available in PMC: 2017 Apr 1.
Published in final edited form as: J Appl Toxicol. 2015 Oct 28;36(4):568–578. doi: 10.1002/jat.3255

Distribution of Single-Wall Carbon Nanotubes in the Xenopus laevis Embryo after Microinjection

Brian D Holt a,†, Joseph H Shawky b,†, Kris Noel Dahl c,d, Lance A Davidson b,*, Mohammad F Islam a,*
PMCID: PMC4943752  NIHMSID: NIHMS759766  PMID: 26510384

Abstract

Single-wall carbon nanotubes (SWCNTs) are advanced materials with the potential for a myriad of diverse applications, including biological technologies and largescale usage with the potential for environmental impacts. SWCNTs have been exposed to developing organisms to determine their effects on embryogenesis, and results have been inconsistent arising, in part, from differing material quality, dispersion status, material size, impurity from catalysts, and stability. For this study, we utilized highly purified SWCNT samples with short, uniform lengths (145 ± 17 nm) well dispersed in solution. To test high exposure doses, we microinjected > 500 μg mL-1 SWCNT concentrations into the well-established embryogenesis model, Xenopus laevis, and determined embryo compatibility and sub-cellular localization during development. SWCNTs localized within cellular progeny of the microinjected cells, but heterogeneously distributed throughout the target-injected tissue. Co-registering unique Raman spectral intensity of SWCNTs with images of fluorescently labelled sub-cellular compartments demonstrated that even at the regions of highest SWCNT concentration, there were no gross alterations to sub-cellular microstructures, including filamentous actin, endoplasmic reticulum and vesicles. Furthermore, SWCNTs did not aggregate or localize to the perinuclear sub-cellular region. Combined, these results suggest that purified and dispersed SWCNTs are not toxic to X. laevis animal cap ectoderm and may be suitable candidate materials for biological applications.

Keywords: Xenopus laevis, carbon nanotube, Raman spectroscopy, nanoparticle toxicity, development, bovine serum albumin, actin

Introduction

Single-wall carbon nanotubes (SWCNTs) possess tremendous potential for new technologies due to their desirable mechanical, electrical and optical properties (Dresselhaus et al. 1996); these properties also offer unique advantages for biological applications (Liu et al. 2009). Because of the potential for developing organisms to be exposed to SWCNTs either intentionally, e.g., using SWCNTs to image and track cells during embryogenesis, or unintentionally, e.g., environmental accumulation of SWCNTs, understanding SWCNT-embryo interactions is crucial for developing SWCNT-based biotechnologies and for minimizing SWCNT ecotoxicity.

Embryogenesis is a critical period of highly coordinated, sophisticated cellular processes, when alterations to the embryonic niche from exogenous molecules may overwhelm inherent embryo defenses and result in developmental defects and/or embryonic lethality (Hamdoun et al. 2007). While substantial effort has been dedicated to studying the implications of numerous nanostructured carbon materials (Campagnolo et al. 2012; Giannaccini et al. 2014) including SWCNTs (Jackson et al. 2013) on development, little focus has been placed on (a) pure, unmodified SWCNT preparations that would be used for biotechnological applications and (b) real-space imaging of SWCNTs co-registered with sub-embryo and sub-cellular structures.

The Xenopus laevis (X. laevis) model is widely used for studying development due to its numerous advantages (Sive et al. 2000). For instance, amphibians are highly sensitive to environmental contaminants and are used as standard models for toxicity screening (Bridges et al. 2002). FETAX (Frog Embryo Teratogenesis Assay – Xenopus) is a standardized assay developed by the U.S.A. Environmental Protection Agency (Bacchetta et al. 2012a; Delgado et al. 2013; Dumont et al. 1983; Hutchison et al. 2014). Some other advantages of the Xenopus embryo model are that fertilization occurs ex vivo at room temperature, the fertilized ovum is about a millimeter in diameter, the embryo develops rapidly from fertilization to gastrulation in approximately 9 hours, and the progressive stages of development can be imaged with a simple stereomicroscope (Sive et al. 2000). X. laevis embryos can be indirectly exposed to materials dispersed in media or directly exposed to materials via microinjection into the embryo. Different tissues can be excised from the embryo using microsurgical manipulations and remain viable which allows for imaging of different surfaces and tissues within the embryo.

SWCNTs possess specific, intense Raman scattering modes, including the G-band at ∼1591 cm-1 and radial breathing modes (RBMs) at ∼100 – 500 cm-1 due to their tubular structure of sp2 hybridized carbon (Dresselhaus et al. 2005). SWCNTs' specific Raman scattering can be confocally imaged with high spatial resolution within biological systems. Raman imaging of SWCNTs offers numerous advantages over other imaging modalities such as bright field absorbance imaging and transmission electron microscopy (TEM). Raman imaging does not require labeling or sample preparation that could alter the specimen, possesses high sensitivity and specificity, has diffraction-limited spatial resolution in the three spatial dimensions and allows for co-registration to fluorescently labeled, specific components.

To determine embryo compatibility and cellular localization of SWCNTs, we microinjected highly purified (∼0.3 wt.% metallic impurities and < 5 wt.% carbonaceous impurities) (Islam et al. 2005; Johnston et al. 2005), short and length fractionated (145 ± 17 nm long) (Holt et al. 2010), primarily individually dispersed (i.e., not in bundles) with bovine serum albumin (BSA) (Holt et al. 2011; Holt et al. 2012a) SWCNTs into X. laevis embryos at the four-cell stage. We analyzed SWCNT-mediated embryo toxicity, imaged the overall distribution of SWCNTs within developing embryos and their sub-cellular localization. SWCNTs did not induce deleterious effects, even at high exposure doses of ∼3 ng per embryo. SWCNTs remained individually dispersed, were distributed throughout the embryo and ultimately were located in the peri-nuclear regions of cells that had internalized the injection volume without altering sub-cellular compartments. These results suggest that well-prepared SWCNTs are not toxic to X. laevis animal cap ectoderm and may be candidate materials for cell biological applications.

Materials and methods

Nanotube dispersions

Single-wall carbon nanotubes were highly purified, length fractionated and well-dispersed as described previously. Briefly, high pressure carbon monoxide conversion synthesis (HiPCO) SWCNTs were purified via wet air burn in the presence of H2O2, acid reflux, magnetic fractionation and annealing (Islam et al. 2005; Johnston et al. 2005). The purified SWCNTs contained < 5 wt.% carbonaceous and ∼0.3 wt.% metallic impurities (Islam et al. 2005) without a change in SWCNT properties (Johnston et al. 2005). SWCNTs were dispersed with deoxycholic acid (Islam et al. 2003) and length fractionated by density gradient ultracentrifugation (Arnold et al. 2006). SWCNTs with lengths of 145 ± 17 nm were selected and the dispersing agent burned off (Holt et al. 2010).

SWCNTs are hydrophobic and interact with each other with an energy of ∼40 kBT/nm (Girifalco et al. 2000; Hough et al. 2004); thus, unmodified SWCNTs are highly bundled in aqueous conditions. Exfoliation into individually dispersed SWCNTs for applications was achieved by sonication of SWCNTs in the presence of a dispersing agent in water, as previously described: 1.0 wt.% bovine serum albumin (BSA) and 0.1 wt.% SWCNTs, subjected to 2 h of probe-tip sonication (Fisher Scientific, Sonic Dismembrator Model 100, 3 mm probe tip) at 6 W (Holt et al. 2011; Holt et al. 2012a). Centrifugation at 21,000×g for 7 min (Holt et al. 2011) was performed to pellet remaining small bundles. Supernatants were decanted and the concentration of SWCNTs was determined by a known absorbance coefficient at 930 nm (Boyer et al. 2013; Holt et al. 2012a; Holt et al. 2012b; Holt et al. 2010). Dispersions were sterilized by UV lamp (UVP Black- Ray, 100 W long-wave ultraviolet lamp, Model B 100 AP/R; UVP, LLC, Upland, CA, USA) for 1 h. Previous description of ultraviolent–visible–near-infrared absorbance, fluorescence and Raman spectroscopies of the BSA-stabilized SWCNTs showing highly resolved van Hove peaks are included elsewhere (Holt et al. 2011; Holt et al. 2012a) and confirm that the supernatants of SWCNTs prepared in this manner contain mostly individualized SWCNTs with some very small bundles (Bachilo et al. 2002; Dresselhaus et al. 1996; Kataura et al. 1999; O'Connell et al. 2002).

Embryos and nanotube microinjection

X. laevis embryos were obtained by standard methods (Kay 1991), fertilized in vitro, dejellied in 2 wt.% cysteine and cultured in (1/3)× Modified Barth's Saline (MBS; 29 mM NaCl, 0.27 mM MgSO4, 0.8 mM NaHCO3, 0.33 mM KCl, 0.11 mM Ca(NO3)2, 0.14 mM CaCl2 and 3.3 mM HEPES) (Sive et al. 2000) at 22 °C. Embryos were microinjected 4× in the animal region at the four-cell stage with a total volume of 2, 4, 8, 16 or 32 nL of 1.0 wt.% BSA suspension or 4 nL of BSA-stabilized SWCNTs at a concentration of 700 μg mL-1 and cultured to tailbud stage (stage 29) or until death. Staging of embryos was performed according to the table of Nieuwkoop and Faber (Nieuwkoop 1967). Embryos were kept in 3% Ficoll solution made in (1/3)× MBS during injection then transferred back to regular (1/3)× MBS after healing (∼30 min).

To isolate animal cap explants, early gastrula (stage 10 – 10.5) embryos were selected and transferred to culture media, Danilchik's For Amy (DFA), containing 53 mM NaCl, 5 mM Na2CO3, 4.5 mM K Gluconate, 32 mM Na Gluconate, 1 mM MgSO4, 1 mM CaCl2 and 1 g of BSA per 1 l, and then buffered to pH 3 with 1 M bicine, for microsurgery. For single cell experiments, animal caps were dissociated by incubation in Ca2+/Mg2+ free DFA for 5 – 10 min and returned back to normal DFA to allow adhesion to the fibronectin-coated substrate (Roche, Mannheim, Germany; 25μg mL-1) before fixation.

Confocal fluorescence imaging

Dissociated cells were fixed in 3.7% formaldehyde / 0.25% glutaraldehyde in phosphate buffed saline (PBS) for 10 – 15 min. F-actin was visualized by incubating fixed cells for 1 – 2 h with BODIPY-FL-Phallacidin (2.5:1000) in PBST (0.1% Triton X). Following phallacidin staining, cells were dehydrated in isopropanol and cleared in Murray's clear (2:1 benzyl benzoate:benzyl alcohol). Single optical sections and Z-stacks of fixed samples were collected using a 63× 1.4 numerical aperture (NA) oil-immersion objective mounted on an inverted compound microscope (DMI6000, Leica Microsystems) equipped with a confocal laser scan head (SP5, Leica Microsystems) using Leica Application Suite Advanced Fluorescence (LASAF, Leica Microsystems) software. Maximum projections and reslicing of z-stacks were carried out with ImageJ (v. 1.38, Wayne Rasband, NIH). Three animal cap explants were subjected to confocal fluorescence imaging and the same explants were separately imaged with confocal Raman spectroscopy. For dissociated cells, at least 40 cells were imaged with confocal fluorescence imaging and then separately imaged with Raman spectroscopy.

Widefield fluorescence imaging

Dissociated cells were fixed in 3.7% formaldehyde / 0.25% glutaraldehyde in PBS for 10 – 15 minutes, permeabilized for 5 min with 0.2% wt.% Triton X-100 and labeled with 4′,6-diamidino-2-phenylindole, dihydrochloride (DAPI) at 0.25 μg mL-1, Oregon Green® 488 phalloidin at 0.165 μM and FM® 4-64 at 25 μM for 30 min (all fluorophores were from Life Technologies). Widefield fluorescence imaging was performed on a Leica DMI 4000B fluorescence and light microscope with a 100× 1.4 NA oil-immersion objective. Over 10 cells were fluorescently imaged and the same cells were separately imaged with Raman spectroscopy.

Raman spectroscopy and imaging

Confocal Raman spectroscopy and imaging was performed on an inverted Raman microscope (inVia Raman microscope, Renishaw) with a 100×, 1.4 NA oil-immersion objective and a 785 nm (1.58 eV) laser of ∼10 mW at the sample plane. Phase contrast imaging was performed on the same microscope including the use of a 0.9 NA condenser. Control of Raman mapping parameters, including 3D position, was performed using WiRE 3.4 software (Renishaw). Different Raman maps were acquired with different step sizes, ranging from 0.5 – 20 μm, depending on the zoom level. Large fields of view were generated by automatically concatenating multiple fields of view together in WiRE software.

The presence of SWCNT G-band in Raman spectra at ∼1591 cm-1 indicates sp2 hybridized carbon structures (Dresselhaus et al. 2005), and we have previously shown that its intensity can be used to determine SWCNT concentration (Boyer et al. 2013; Holt et al. 2011; Holt et al. 2012a; Holt et al. 2012b). The intensity ratio ID:IG between the SWCNT D-band, which characterizes the sp3-hybridized carbon, and the G-band indicates damage or structural defects in SWCNTs. For SWCNTs prepared via HiPCO synthesis, their diameters are ∼1 nm, and when subjected to Raman spectroscopy with a 785 nm (1.58 eV) laser, RBMs < 250 cm-1 arise from individualized, that is not bundled, SWCNTs whereas RBMs > 250 cm-1 arise from bundles of SWCNTs (Dresselhaus et al. 2005; Heller et al. 2004). This is because the SWCNT (10,2) chirality comes into resonance with the 785 nm laser when it is within bundles of SWCNTs but not when it remains individually dispersed (Heller et al. 2004). Also, real space SWCNT near-infrared (NIR) fluorescence was also detected in the Raman spectra from ∼2100 – 3000 cm-1 which only arises from individualized SWCNTs (O'Connell et al. 2002). We have previously used these Raman spectral features to determine SWCNT dispersion state within cells (Holt et al. 2012a). A Raman spectrum of the initial SWCNT dispersion, fluorescence spectroscopy of SWCNTs, a summary of the SWCNT diameters, and further discussion on using the RBMs to determine SWCNT bundling is included in Supporting Information, Fig. S1.

Therefore, RBMs can be used to probe local SWCNT dispersion status in cells (Holt et al. 2012a). For mapping experiments, spectra were acquired at Raman shifts of 1188 – 1696 and 197 – 802 cm-1 for G-band and RBM scans, respectfully. Analysis of Raman mapping experiments was performed using in-house MATLAB® (The MathWorks, Inc.) code: G-band and RBM signal above baseline (having a signal-to-noise ratio >3) was used to color-code intensity maps. For G-band, the code was especially designed to accurately determine G-band signal while differentiating it from other signals from the explants at ∼1580 cm-1 and ∼1600 cm-1, in addition to determining non-uniform backgrounds.

Fluorescence/SWCNT overlays

Confocal fluorescence Z-stacks and widefield fluorescence images were acquired on separate imaging systems than Raman maps. To co-register fluorescence and Raman signals, the same field of view was imaged on both systems. Then, as indicated, confocal or widefield fluorescence images were overlaid with confocal Raman images by matching the fluorescence signals (confocal) or phase image (widefield) with the phase image acquired on the Raman system. Images were adjusted to be the same size and overlays were generated using LASAF software.

For Z-stack overlays of confocal fluorescence and confocal Raman, more processing needed to be performed as Raman Z-stacks were sub-sampled compared to fluorescence: Raman Z-step size was 2 μm was fluorescence was 0.2 μm. Therefore, Raman Z-stacks were interpolated (spline interpolation in MATLAB) to generate the same number of images as fluorescence. Alignment of Z-positions was based on where confocal F-actin intensity began and where phase contrast (on the Raman system) just came into focus. While theoretically lipid or other biological molecule peaks could be used to determine the Z-position on the Raman system, practically there was not enough signal to discriminate these peaks. While phase contrast is not a confocal imaging modality, similar processing was performed to it to create Z-stack, and phase intensity indeed varied with Z-position.

Results

Single-wall carbon nanotubes are not toxic

To maximize direct exposure of SWCNTs to X. laevis, we microinjected suspensions of SWCNTs into the animal hemisphere of all four blastomeres at the four-cell stage. To determine appropriate injection volumes of SWCNT suspensions, we first microinjected a series of volumes (2, 4, 8, 16 and 32 nL) of the dispersing agent BSA at its concentration used in the dispersion procedure, 1.0 wt.%, to determine the non-toxic, upper-limit injection volume. Microinjection of 4 nL of 1.0 wt.% BSA suspension resulted in normal development to tailbud stage (stage 29) while volumes > 4 nL were toxic and inhibited proper development (Table 1). Embryos injected with 8 nL developed defects including stalled gastrulation and/or shortened axes; injections of 16 nL induced early lethal defects including failure of blastomeres to cleave properly. Hence, all subsequent experiments were performed using an injection volume of 4 nL.

Table 1.

Toxicity of BSA and BSA-dispersed SWCNTs in X. laevis embryos.

Group 8-cell St. 11 St. 14 St. 29
Control 20/20 healthy 20/20 healthy 20/20 healthy 20/20 healthy
1.0 wt.% BSA 4 nL 20/20 healthy 1/20 stalled gastrulation 20/20 healthy 20/20 healthy
8 nL 20/20 healthy 6/20 stalled gastrulation 2/20 exogastrulae* 14/18 shortened axes
16 nL 20/20 healthy 12/20 stalled gastrulation 15/20 exogastrulae* 5/5 shortened axes
SWCNTs–BSA 4 nL of 700 μg mL-1 20/20 healthy 20/20 healthy 20/20 healthy 20/20 healthy
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*

Exogastrulae are embryos that fail to gastrulate, specifically in which the endomesoderm and yolk plug are extruded outwards. These embryos were removed from culture upon observation.

To determine whether microinjections of SWCNTs into the animal hemisphere of the X. laevis embryo induced any developmental defects, we injected the non-toxic upper-limit volume of 4 nL of highly concentrated (700 μg mL-1) SWCNTs into prospective animal cap tissue cells at the 4-cell stage. The animal cap is a dual layer tissue composed of deep mesenchymal cells with an overlying epithelium that is the precursor to the skin and nervous system of the tadpole. This tissue can be precisely targeted by microinjection and is easily visualized in vivo or in vitro by microsurgically excising it from the rest of the embryo followed by culturing on fibronectin. Tissue movements in the animal cap, including radial intercalation and outward spreading are well characterized. Therefore, variations from native behaviors due to exogenous influences such as SWCNT injection would be easily detected. Injections of SWCNTs into these tissues yielded no deleterious effects on development and were nontoxic (Table 1). Embryos successfully developed through tailbud stage (stage 29), demonstrating that microinjection of ∼3 ng of SWCNTs into embryos did not affect X. laevis development.

Nanotubes are distributed throughout the animal cap ectoderm

To determine the localization of SWCNTs within developing embryos, microinjected X. laevis embryos were allowed to develop to gastrula stage (stage 10 – 10.5) when animal cap ectoderm tissues were microsurgically isolated, fixed and mounted for imaging. Animal cap ectoderm was imaged using high resolution phase contrast microscopy, and the spatial distribution and local concentration of SWCNTs were imaged using Raman spectroscopy. We imaged 10 separate animal caps, and the results were consistent between samples. Co-registration of embryo features with SWCNT signal revealed that SWCNTs were heterogeneously distributed throughout the animal cap (Fig. 1A). Some regions possessed high SWCNT concentrations (> 100 μg mL-1) which were approximately similar to the concentration of the injection (700 μg mL-1), while many regions throughout the animal cap tissue contained less concentrated SWCNTs (< 25 μg mL-1). Higher spatial resolution heat maps of the distribution and concentration of SWCNTs revealed centers of highly concentrated SWCNTs away from which concentration decreased (Fig. 1B). There were no observable alterations to embryonic tissue structures in regions with SWCNTs, even for those with high concentrations.

Figure 1.

Figure 1

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Raman imaging of the localization of BSA-dispersed SWCNTs in explanted animal cap ectoderm tissues. (A) Zoomed out views of animal cap ectoderm reveal that SWCNTs are distributed throughout the explants, with some regions having high concentrations. Scale bars are 100 μm. (B) Images at increasing zoom and spatial resolution show detailed localization of SWCNTs. Scale bars are 10 μm. (C) Raman spectra from the positions indicated demonstrating the RBMs (200 – 250 cm-1), D-band (∼1300 cm-1), G-band (∼1591 cm-1), G'-band (∼2600 cm-1) and NIR fluorescence (2100 – 3000 cm-1), in addition to numerous other peaks arising from molecules of the animal cap ectoderm. The presence of RBMs < 250 cm-1 and fluorescence confirm that SWCNTs remained individually dispersed.

The dispersion state of SWCNTs, or in other words if SWCNTs remained individually dispersed or became bundled, is important for the physical properties of SWCNTs (Dresselhaus et al. 1996; Hersam 2008; Jariwala et al. 2013; O'Connell et al. 2002) and biological interfacial properties (Battigelli et al. 2013; Bilalis et al. 2014; Heister et al. 2013). Raman spectra of SWCNTs were used to determine the dispersion state of SWCNTs. Spectra contained numerous peaks arising from the animal cap itself; nonetheless, peaks that are specific for SWCNTs were readily detected, including the G-band at ∼1591 cm-1 and RBMs between 200 – 275 cm-1 (Fig. 1C). The relatively high ratio of RBM intensity < 250 cm-1 to RBM intensity > 250 cm-1, along with the presence of NIR fluorescence of SWCNTs, manifesting as a broad peak within 2100 – 3000 cm-1, confirmed the presence of individually dispersed SWCNTs and that SWCNTs did not aggregate into large bundles.

Nanotubes are distributed in three-dimensions

To determine the three dimensional (3D) spatial distribution of SWCNTs within X. laevis embryos and whether microinjection of SWCNTs induced disruption of the earliest manifestation of multi-laminate tissue structure, we performed confocal Raman imaging of the intensity of the G-band of SWCNTs co-registered with confocal fluorescence imaging of rhodamine phalloidin that labeled filamentous-actin (F-actin) structures. We imaged F-actin because it is a highly prevalent protein, plays crucial roles in numerous cellular processes including division, motility, endocytosis etc. (Pollard et al. 2008) and could be used to delineate individual cells.

Confocal maximum Z-projected stacks of animal caps microinjected with SWCNTs showed normal F-actin structures (Fig. 2) and were consistent between samples. SWCNTs were distributed throughout the animal cap ectoderm in 3D, and SWCNTs neither co-localized with F-actin structures nor induced alterations to F-actin structures.

Figure 2.

Figure 2

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3D spatial localization of SWCNTs within animal cap ectoderm. Images are maximum projections in Z-direction, Y-direction (bottom), and X-direction (right). Note that all Y- and X-direction projections are anisotropically stretched by 5× as indicated to increase visual clarity. Confocal fluorescence Z-stacks of F-actin were co-registered to confocal Z-stacks of the Raman G-band intensity of SWCNTs. Z-stacks of phase contrast images were also acquired and are included in a separate overlay for clarity. The cyan dashed boxes indicate the spatial regions in which Raman spectra were acquired. Note that SWCNT signal is not co-registered with altered F-actin structures or highly phase-dense regions.

Nanotubes are localized within cells that internalized the injection volume

To further investigate how SWCNTs were distributed throughout the X. laevis animal cap, we co-injected SWCNTs with 0.5 ng of rhodamine-conjugated 10 kDa dextran per blastomere at the four cell stage. Labeled dextrans microinjected into embryos do not transfer from cell to cell and localize to the cytoplasm of injected cells where they can be used for lineage tracing (Gimlich et al. 1985). Therefore, the co-distribution of SWCNTs and fluorescently-labeled dextrans could demonstrate whether SWCNTs remained localized in the cytoplasm of injected cells.

Due to the opacity of X. laevis embryos and the need to accurately co-register Raman images with fluorescence images, we performed single cell layer imaging rather than imaging the entire, intact animal cap explants. To enable single cell imaging, we dissociated animal caps, allowed the individualized cells to adhere to the substrate, then fixed and labeled all cells with BODIPY® FL phallacid into label F-actin.

Confocal maximum Z-projections of F-actin and dextrans clearly indicated that some cells inherited fluorescently labeled dextrans while other cells did not (absence of fluorescence from dextrans but presence of fluorescence from F-actin structures) (Fig. 3 and Supporting Information, Fig. S2). Confocal Raman imaging of the same field of view reveal SWCNTs in cells that are fluorescently labeled with dextrans. These results were consistent across all samples. Therefore, SWCNTs were incorporated into and distributed throughout the progeny of the cells exposed to the injection bolus, like the fluorescently labeled dextrans.

Figure 3.

Figure 3

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Cellular distribution of SWCNTs. Fluorescently labeled dextrans, which are known to localize within the progeny of cells that internalized the injection volume, were also microinjected with SWCNTs. Animal caps were dissociated to allow for individual cell imaging and unambiguous determination of co-localization. Confocal fluorescence images of F-actin (present in all cells) and dextrans (present in the progeny of cells that internalized the injection volume) were co-registered to confocal Raman images of SWCNT G-band and RBM intensities. The co-localization of SWCNTs and dextrans confirms that SWCNTs are present in cells that internalize the bolus but not in neighboring cells. Intense RBM signal < 250 cm-1 demonstrates that SWCNTs remain individualized (i.e., do not bundle). SNR is an abbreviation for signal-to-noise ratio. A representative extended scan shows the peaks characteristic of SWCNTs and also SWCNT NIR fluorescence (∼2100–3000 cm-1).

To determine the dispersion state of SWCNTs, we acquired Raman spectra of the RBMs of SWCNTs and plotted heat maps of the intensity of RBMs that arise from individualized SWCNTs (Fig. 3 and Supporting Information, Fig. S2). Like the injected dispersion, SWCNTs possessed significant RBM intensity from individualized SWCNTs, and a representative Raman spectrum clearly showed high intensity of RBMs from individually dispersed SWCNTs (RBM intensity < 250 cm-1) along with broad NIR fluorescence of SWCNTs (∼2100 – ∼3000 cm-1), confirming that SWCNTs remained individually dispersed and were not bundled. Additionally, the ID:IG was 0.05 ± 0.02 for the intracellular SWCNTs, which is similar to that of SWCNTs before microinjection of 0.03 ± 0.02.

Nanotubes are localized in peri-nuclear regions and do not alter sub-cellular compartments

To determine the sub-cellular localization of SWCNTs and potential alterations to sub-cellular compartments, we fluorescently labeled important cellular compartments and imaged individual cells. To do so, we dissociated animal cap ectoderm tissues, allowed the individualized cells to adhere to the fibronectin-coated substrate, then fixed and labeled the cells with DAPI to label DNA, Oregon Green phalloidin to label F-actin and FM 4-64 to label endoplasmic reticulum (ER) and vesicles.

Fluorescence imaging of cellular compartments consistently revealed no gross change to their localization or morphology (Fig. 4 and Supporting Information, Fig. S3). Co-registering fluorescence images of cellular components and the Raman image of SWCNT concentration revealed high concentrations of SWCNTs that were not co-localized with sub-cellular structures but were primarily located in the peri-nuclear regions. SWCNTs did not alter the localization of the sub-cellular compartments, even though some regions had concentrations > 100 μg mL-1. For selected regions of high SWCNT concentrations, higher spatial resolution (smaller step size between spectral acquisitions) Raman imaging further confirmed that SWCNTs were located in the peri-nuclear regions without alterations to cellular compartments (insets of Fig. 4).

Figure 4.

Figure 4

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SWCNT sub-cellular localization. Fluorescence images of dissociated animal cap ectoderm tissues co-registered to SWCNT Raman G-band intensity images demonstrate that SWCNTs are located in peri-nuclear regions. Zoomed-in, high spatial resolution Raman images further demonstrate that SWCNTs do not alter F-actin or ER structures.

To obtain high spatial resolution images of SWCNT dispersion status, Raman spectra of the RBMs of SWCNTs were acquired. Imaging revealed substantial intensity of RBMs < 250 cm-1, demonstrating that SWCNTs remained well dispersed (i.e., did not aggregate) within cells although their local concentration remained high (Supporting Information, Fig. S3).

Discussion

Singl-wall carbon nanotubes are not toxic

Several studies have found embryonic defects induced by nanostructured carbon materials, including nanodiamonds (Marcon et al. 2010; Vankayala et al. 2014; Wierzbicki et al. 2013), paracrystalline carbon powder (Bacchetta et al. 2012b; Cheng et al. 2007; Jackson et al. 2012; Pietroiusti et al. 2011), fullerenes (Kim et al. 2012; Murugesan et al. 2007; Vankayala et al. 2014; Wierzbicki et al. 2013), graphite (Chen et al. 2012; Murugesan et al. 2007; Wierzbicki et al. 2013), multi-wall carbon nanotubes (Chen et al. 2012; Cheng et al. 2009; Cheng et al. 2012; Hougaard et al. 2013; Huang et al. 2014; Kim et al. 2012; Liu et al. 2014; Murugesan et al. 2007; Nouara et al. 2013; Wierzbicki et al. 2013) and SWCNTs (Campagnolo et al. 2013; Chen et al. 2013; Cheng et al. 2007; Cheng et al. 2011; Huang et al. 2014; Leeuw et al. 2007; Pietroiusti et al. 2011; Roman et al. 2013). However, relatively few studies have used the X. laevis model, even though it is especially advantageous to examine potential cytotoxic effects of nanomaterials because it highly sensitive to environmental contaminants and provides an in vivo environment in which cells are undergoing cell division, differentiation, migration, and vesicular transport all while maintaining adhesions with neighboring cells and extracellular matrix (Bridges et al. 2002).

Prior studies using X. laevis embryos found nanostructured carbon materials could be toxic. For example, microinjected, nearly spherical ∼4 nm diameter nanodiamonds had toxicity that was highly dependent on nanodiamond interfacial properties, as carboxyl functionalization resulted in embryotoxicity and teratogenicity but hydroxyl functionalization was not toxic, even for a 10 nL microinjection of 2 mg mL-1 nanodiamonds (Marcon et al. 2010). Other studies exposed X. laevis to nanostructured carbon materials in suspension. A series of studies from the same group determined that undispersed carbon nanotubes were cytotoxic but not genotoxic (Mouchet et al. 2011; Mouchet et al. 2010; Mouchet et al. 2008; Saria et al. 2014). Double-wall carbon nanotubes accumulated in the gills and intestines, resulting in growth inhibition and embryotoxicity but minimal genotoxicity (Mouchet et al. 2011; Mouchet et al. 2008). Similar results were obtained for multi-wall carbon nanotubes, as they inhibited growth, increased oxidative stress and induced DNA damage (Mouchet et al. 2010; Saria et al. 2014). Other groups also demonstrated that multi-wall carbon nanotubes were embryotoxic at exposure doses of 50 μg mL-1 but were not genotoxic (Bourdiol et al. 2013). Amorphous carbon spheres of diameters of 28.5 ± 14.3 nm also led to toxicity at an exposure dose of 100 μg mL-1, and the nanoparticles preferentially accumulated in the stomach and gut (Bacchetta et al. 2012b).

Many of these studies utilized materials that were either not highly purified or not individually dispersed. Many nanostructured carbon materials are synthesized with toxic catalysts, and syntheses typically also produce carbonaceous impurities. Without extensive purification, material/embryo interactions may be dominated by these minority components. Further, as nanostructured carbon materials are hydrophobic, this can lead to aggregation of not well dispersed materials in biological environments, complicating material/embryo interactions. While studies of as-produced or minimally processed materials are crucial for understanding the toxicity of these materials that, for example, may be used in industrial applications, they are not directly applicable to highly purified, well-dispersed materials that, for example, may ultimately be utilized for biological applications.

Therefore, in this study we investigated ultrapure (∼0.3 wt.% metal and < 5 wt.% carbonaceous impurities), length selected (145 ± 17 nm), individually dispersed SWCNTs (∼1 nm diameter, individual SWCNTs) (Holt et al. 2011; Holt et al. 2012a; Holt et al. 2010), as this preparation has potential for use in biotechnological. SWCNTs were purified to remove virtually all toxic catalyst, length fractioned to avoid altered cellular interactions due to lengths varying over a couple orders of magnitude and individually dispersed to remove bundles that have larger diameters that could affect interactions with embryo structures and reduce SWCNT properties.

These highly processed SWCNTs possessed no toxicity to X. laevis embryos up to ∼2.8 ng of SWCNTs delivered via 4 nL microinjection directly into their cytoplasm.4 nL was an upper limit on injection volume because the dispersing agent control solution of 1.0 wt.% BSA caused toxicity for injections > 4 nL. We attribute the toxicity of higher injection volumes to the surfactant-like nature of BSA (Curry et al. 1999) that could solubilize embryo proteins and membranes and the osmotic pressure of the ultrapure water that could substantially affect local osmolarity and pH, as observed previously (Dubertret et al. 2002). ∼700 μg mL-1 was a reasonable upper limit on SWCNT concentration since concentrations > ∼3 mg mL-1 alter SWCNT state (Hough et al. 2006; Puech et al. 2011). Therefore, microinjecting ∼3 ng of SWCNTs was an effective upper limit for our system, and at this maximum exposure dose, there was no SWCNT toxicity.

We attribute the lack of toxicity of our SWCNTs due to the extensive materials preparation. By removing virtually all metal catalyst that remained from SWCNT synthesis (Islam et al. 2005; Johnston et al. 2005), we avoided metal impurity toxicity that is unavoidable for unpurified SWCNTs (Kagan et al. 2006). Length selecting SWCNTs to be 145 ± 17 nm long enabled endocytosis and exocytosis (Becker et al. 2007). Individually dispersed SWCNTs have been shown to be more cytocompatible than bundles, enabling organism and cellular recovery from SWCNT exposure and allowing for long-term cellular labeling without altering cellular state or function (Heller et al. 2005; Holt et al. 2012a; Jin et al. 2009; Jin et al. 2008; Kolosnjaj-Tabi et al. 2010; Liu et al. 2013; Ruggiero et al. 2010).

Injected nanotubes distribute throughout the targeted animal cap ectoderm

Unlike many studies that solely relied on the presence of dark regions in bright field microscopy or relatively large aggregates of SWCNTs in TEM to identify the spatial localization of SWCNTs, we utilized SWCNTs' optical property of unique Raman scattering to specifically and sensitively non-invasively image SWCNT distribution without requiring extensive sample preparation that could potentially alter SWCNT sub-embryo localization. While highly optically absorbing regions of bright field images may allow for rapid, accessible imaging, dark regions are not necessarily SWCNTs and light regions are not necessarily SWCNT-free, and these exceptions have been experimentally reported for X. laevis embryos (Mouchet et al. 2011; Mouchet et al. 2008). TEM of multi-wall carbon nanotubes in X. laevis larvae required careful staining to generate images of proper contrast and sectioning samples with an ultramicrotome which may have altered their distribution within the biological matrix (Bourdiol et al. 2013). Here, we acquired Raman spectra in a point-by-point manner and generated heat maps of SWCNTs' unique, intense G-band (Dresselhaus et al. 2005). Additionally, by utilizing a previously established calibration curve (Holt et al. 2012a), we were able to quantitatively determine the local concentration of SWCNTs.

Utilizing the high sensitivity, we determined that SWCNTs were distributed throughout the cells of the animal cap in a somewhat heterogeneous manner. Many SWCNT-containing regions had concentrations ∼10× lower than the injection dose while a few regions remained highly concentrated, similar to the injection dose concentration. These results suggest that the cells that were initially injected subsequently proliferated and distributed SWCNTs throughout their daughter cells. Overall, the SWCNTs were inert and did not alter embryonic development or structures of the animal cap ectoderm.

Nanotubes are distributed in 3D

While confocal fluorescence and Raman imaging enabled the generation of 3D stacks of co-registered images, there were some limitations. Raman imaging was performed in the X–Y plane over a Z-height of 10 μm with a 2 μm Z-direction step size. Since the F-actin fluorescence was acquired with a 0.2 μm step size, the Raman and phase contrast images were interpolated to create Z-stacks of the same number of images to enable overlaying. Additionally, confocal fluorescence and confocal Raman imaging were performed using different imaging modalities; therefore, there was some inherent uncertainty in the co-registration of spatial localization between the two imaging systems. Finally, phase contrast images, which are not confocal, were acquired at each Raman Z-step and used to align the Raman spatial position with the fluorescence position. Nonetheless, we were able to generate Z-stacks that demonstrated SWCNTs' 3D localization within the animal cap ectoderm of X. laevis embryos.

Nanotubesare retained by cell lineages that were injected

Fluorescently labeled dextrans have been used as cellular lineage tracers, as microinjections are uniformly distributed throughout the cytoplasm of the progeny of the microinjected cells (Gimlich et al. 1985). Therefore, fluorescently labeled dextrans can be used to image the cellular progeny of the microinjected cells. By dissociating animal caps, we were able to accurately co-register SWCNT cellular localization with fluorescently labeled dextrans and probe sub-cellular SWCNT distribution. Raman imaging demonstrated that SWCNTs were present within cells that also contained the fluorescently labeled dextrans but were not present in progeny from cells that did not receive the microinjection of SWCNTs, as expected (Fig. 3 and Supporting Information Fig. S2). This distribution of SWCNTs is in agreement with previous results that microinjections of well dispersed quantum dots into X. laevis embryos were cell autonomous, that is the quantum dots were retained within the progeny of the microinjected cell (Dubertret et al. 2002).

As indicated for quantum dots, stability of the nanoparticle dispersion is important for cytocompatibility and ultimate use for biotechnological applications (Dubertret et al. 2002). We have previously shown that the BSA-dispersed SWCNTs used in this study were stable upon dilution in ultrapure water (Holt et al. 2011) and cell culture media (Boyer et al. 2013; Holt et al. 2012b). Therefore, SWCNTs should have remained well dispersed after injection into X. laevis, and indeed, Raman spectroscopy confirmed that SWCNTs remained well dispersed within X. laevis. Thus, we suggest that since SWCNTs remained individualized, this facilitated their distribution throughout the cellular progeny that recapitulated the distributions of the simultaneously microinjected fluorescently labeled dextrans.

Nanotubes are sub-cellularly localized in peri-nuclear regions and do not alter sub-cellular compartments

There have been reports in the literature by our group (Holt et al. 2015; Holt et al. 2012c; Holt et al. 2010; Shams et al. 2014; Yaron et al. 2011) and others (Dulinska-Molak et al. 2014; Kaiser et al. 2008; Rodriguez-Fernandez et al. 2012; Sargent et al. 2012; Villegas et al. 2014; Zhang et al. 2011) that have shown that high levels of cellular exposure to SWCNTs can alter sub-cellular components, including F-actin structures. Even though SWCNTs were present in X. laevis embryos at high local concentrations (up to ∼2 mg mL-1), there were no gross alterations to sub-cellular morphologies, including F-actin and ER. Instead, SWCNTs were located in peri-nuclear regions, as we have previously observed for these SWCNTs extracellularly exposed to and internalized into human and murine 2D cell culture (Boyer et al. 2013; Holt et al. 2011; Holt et al. 2012a; Holt et al. 2015; Holt et al. 2012b). Instead of SWCNTs aggregating into bundles which have been shown to be toxic (Kaiser et al. 2008; Liu et al. 2010) within cells due to strong van der Waals interaction energy between bare SWCNTs (Girifalco et al. 2000; Hough et al. 2004), they remained individually dispersed, which has been shown to play important roles in cellular and organism recovery from SWCNT exposure (Holt et al. 2012a; Holt et al. 2015; Jin et al. 2009; Kolosnjaj-Tabi et al. 2010; Liu et al. 2013; Ruggiero et al. 2010).

Extension to potential applications

Understanding how SWCNTs interact with developing embryos also yields insight into SWCNT interactions with stem cells, since injections occur at the 4-cell stage into pluripotent cells. Since embryos exposed to SWCNTs successfully developed throughout the length of the experiment, tailbud stage (stage 29), SWCNTs did not substantially alter pluripotent cell proliferation or differentiation.

These results indicate that purified, short, well dispersed SWCNTs have promise for many potential developmental and biotechnological applications. For example, the unique optical properties of SWCNTs exist in the NIR “biological window” (Hong et al. 2012; Welsher et al. 2011) which may allow for real-time imaging and/or photothermal strategies to probe development, or SWCNTs' high surface-area-to-volume ratio may allow for targeted, bioactive molecule delivery.

Conclusions

Injections of highly purified, length-selected, SWCNTs well dispersed with BSA into the animal region of X. laevis embryos were not toxic. SWCNTs were somewhat heterogeneously distributed throughout the animal cap ectoderm, specifically in the progeny of the cells that internalized the injection bolus. Confocal fluorescence imaging of cellular compartments co-registered to confocal Raman imaging of SWCNT G-band intensity revealed that SWCNTs were located in peri-nuclear regions, without grossly altering sub-cellular structures. SWCNTs remained well-dispersed in embryos, preserving properties of individualized SWCNTs and preventing deleterious effects of bundles. Our results suggest that purified, short, dispersed SWCNTs, after additional studies, may be candidate materials for diverse biological applications.

Supplementary Material

Figure S1. Optical characterization of SWCNTs. (A) Fluorescence spectroscopy of SWCNTs acquired using a Nanolog Spectrofluorometer with a liquid-nitrogen-cooled Symphony InGaAs 1700 detector (Horiba Jobin Yvon). The heatmap intensities are normalized to the maximum signal. Nanosizer (Horiba Jobin Yvon) was used to determine SWCNT chiralities (n,m) using a Voigt 2D model. (B) The diameter (abbreviated Dia) of the SWCNT chiralities detected using fluorescence spectroscopy. (C) Raman spectrum of the SWCNTs before microinjection. For a 785 nm laser (used for this study), the RBM peak for the SWCNT (10,2) chirality (at 266 cm-1) is only in resonance when the (10,2) chirality is in a bundle of SWCNTs. Thus, it can be used to determine if SWCNTs become bundled.

Figure S2. Additional field of view demonstrating that SWCNTs were co-localized in the cells that contained fluorescently labeled dextrans which were the cellular progeny of the cells that internalized the injection volume. SNR is an abbreviation for signal-to-noise ratio.

Figure S3. Additional field of view showing the sub-cellular localization of SWCNTs. Maps of the intensity of the G-band and RBMs (< 250 cm-1) do not co-register with F-actin or ER and vesicles and confirm peri-nuclear sub-cellular localization. SNR is an abbreviation for signal-to-noise ratio.

Acknowledgments

This work was supported by: the National Science Foundation (CMMI-1335417 [M.F.I.]; CAREER IOS-0845775 and CMMI-1100515 [L.A.D.]; CMMI-1300476 [K.N.D.]); and the National Institutes of Health (R01 HD044750 [L.A.D.]). Any opinions, findings, and conclusions or recommendations expressed in this material are those of the authors and do not necessarily reflect the views of the NSF or the NIH.

Footnotes

Conflict of interest: The authors did not report any conflict of interest.

Supporting information: Additional supporting information may be found in the online version of this article at the publisher's web-site.

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Supplementary Materials

Figure S1. Optical characterization of SWCNTs. (A) Fluorescence spectroscopy of SWCNTs acquired using a Nanolog Spectrofluorometer with a liquid-nitrogen-cooled Symphony InGaAs 1700 detector (Horiba Jobin Yvon). The heatmap intensities are normalized to the maximum signal. Nanosizer (Horiba Jobin Yvon) was used to determine SWCNT chiralities (n,m) using a Voigt 2D model. (B) The diameter (abbreviated Dia) of the SWCNT chiralities detected using fluorescence spectroscopy. (C) Raman spectrum of the SWCNTs before microinjection. For a 785 nm laser (used for this study), the RBM peak for the SWCNT (10,2) chirality (at 266 cm-1) is only in resonance when the (10,2) chirality is in a bundle of SWCNTs. Thus, it can be used to determine if SWCNTs become bundled.

Figure S2. Additional field of view demonstrating that SWCNTs were co-localized in the cells that contained fluorescently labeled dextrans which were the cellular progeny of the cells that internalized the injection volume. SNR is an abbreviation for signal-to-noise ratio.

Figure S3. Additional field of view showing the sub-cellular localization of SWCNTs. Maps of the intensity of the G-band and RBMs (< 250 cm-1) do not co-register with F-actin or ER and vesicles and confirm peri-nuclear sub-cellular localization. SNR is an abbreviation for signal-to-noise ratio.

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