Chapter Summary

This chapter highlights some recently published outstanding researches that have pushed further the boundary of photoluminescent carbon nanomaterials applications to biological systems and have brought invaluable insights into cellular and organism dynamics. From our perspective, there is a need for nanoparticle‐based methodologies that allow researchers to access the spatiotemporal dynamics inherent to a wide variety of biological processes.

Two carbon allotropes have recently shown remarkable advances in neurosciences: fluorescent nanodiamonds (FND) [1], and single‐walled carbon nanotube (SWCNT) [2]. This chapter highlights three examples of recent archetypical achievements based on the tracking of FND or SWCNT while being aware that the list is not exhaustive. Single‐particle tracking (SPT) of FND was used to monitor the endosomal transport inside hippocampal neurons dissociated from mouse embryos and the author used this nanoparticle‐based technique to unravel transport defects in mouse models of brain diseases. Single particle tracking of SWCNT was used to study the nanoscale organization and rheological properties of the extracellular space in acute slices of mouse brain. Altogether, this chapter reviews a decade of research related to FND and SWCNT tracking in cells (such as cancer cell lines and neurons) and organisms (such as zebrafish, drosophila embryos, C‐elegans and mice).

6.1 Introduction

Many carbon allotropes nanoparticles are currently used in biology and biomedical applications, including fluorescent diamond nanocrystals, single‐walled carbon nanotubes, carbon dots, reduced graphene oxide, and fullerenes. Each of them often have specific applications. For example, fullerenes are mostly used for drug delivery in tumor therapy and for theranostic applications [3], whereas reduced graphene oxide is mainly used as a biomolecule sensor owing to its ability to strongly quench dye fluorescence [4]. Carbon dots was used for cellular imaging via passive internalization in cell lines, although no appealing dynamic biological experiments were conducted [5]. The two carbon nanomaterial allotropes that show great potentials are FND and SWCNT. They were shown to be used individually (like in extracellular or intracellular single particle tracking) or in clusters (like in tracking cellular system). This chapter starts with tracking cells using FND, followed by a section of single‐particle tracking of FND inside living organism and inside living cancer cells and neurons from mouse embryos. The last section deals with the use of SWCNT for the study of the extracellular space nanoscale dimensions and local viscosity in live brain tissue.

6.2 Tracking Cells in Organisms with Fluorescent Nanodiamonds

Many studies have shown that acid‐treated high‐pressure high‐temperature (HPHT) synthesized 100 nm‐sized (and smaller) nanodiamonds (ND) can be efficiently internalized inside living cells and organisms. This spontaneous uptake happens in diverse cell types, including cancer cells [6] and primary neurons [1], following the common pathway of receptor mediated endocytosis [7]. NDs are then trapped in endocytotic compartments at different stages of their evolution, down to the lysosomes [1, 7]. ND cellular internalization has very distinct characteristics, with (i) almost no exocytosis even during cell proliferation [8], and (ii) a very low induced toxicity on a few days scale as assessed from cell survival, morphogenesis [9], functions [10], and genotoxicity [11] measurements.

ND can be made fluorescent in the near‐infrared spectral region (600–750 nm wavelength range) by creating a nitrogen‐vacancy (NV) color center within the diamond lattice, which requires the association of naturally present nitrogen (N) impurities (≈100 ppm abundance in HPHT diamond) with vacancy (V) induced by high energy particle irradiation [12, 13]. NV‐centers in diamond have remarkable properties [14]: (i) a perfect photostability (no bleaching and no blinking) and (ii) an optically detectable magnetic resonance (ODMR) of the electronic spin of its negatively charged form (NV⁻ ). The latter offers the possibility to modulate the fluorescence signal of FND with a variable external magnetic field [15]. Furthermore, when NV centers are embedded in diamond nanoparticles of size much smaller than the optical excitation wavelength, their radiative lifetime substantially increases by a dielectric screening effect [16]. For example, the fluorescent lifetime of ≈100 nm sized FND is ≈30–40 ns (compared to 12.7 ns in bulk). Such value is one order of magnitude longer than tissue autofluorescence lifetime (≈1–2 ns). This offers a simple way to enhance the FND signal‐to‐background ratio by combining pulsed laser excitation with time‐gated detection [6]. Altogether, FND gathers a unique set of properties that makes it particularly suited tool for stem cell and regenerative medicine researches, because these fields require that cell proliferation and differentiation be tracked on long‐term scale [8].

Stem cells are nondifferentiated cells that can differentiate in specialized cell types and can go through numerous cycles of cell division. For these reasons, therapies based on stem cell injection or transplantation have been developed to treat a wide range of high‐prevalence diseases, including cancers and neurodegenerative diseases. To optimize stem cell therapy, it is crucial to track stem cells fate in the organism where they have been transplanted. Several strategies are used to label stem cells in vivo [17] , including (i) the overexpression of a reporter gene and the subsequent labeling of the corresponding protein with a molecular probe associated to either a radiotracer (then imaged by positron emission tomography or single‐photon emission computed tomography), or a fluorescent dye; and (ii) internalization of magnetic or fluorescent nanoparticles into stem cells that allow them to be traced by magnetic resonance imaging or photonic imaging. The genetic transformation of cells and their subsequent labeling is a powerful but complex approach with cell viability issues and risks of mutagenesis. Labeling with nanoparticles can offer long signal persistence, which represents a significant advantage. In this category, FND has been successfully used to label lung stem cells (LSCs) and track their engraftment and regenerative capability in mice [18]. FND‐labeled LSCs were injected intravenously into lung‐injured mice, after having checked that FND labeling does not suppress the cells properties of self‐renewal and differentiation into pneumocytes. Mice were sacrificed at different time points and organs were collected and sectioned to evaluate the capability of LSC to stimulate lung regeneration by its engraftment and differentiation. Raster scanning time‐gated fluorescence (TGFluo) confocal microscopy [6] and fluorescence lifetime imaging (FLIM) allowed unambiguous identification of FND‐labeled LSCs despite tissue immunostaining and autofluorescence. Figure 6.1 displays lung section immunochemical staining, FLIM and TGFluo images of a mouse sacrificed seven days after LSCs injection. Immunostaining revealed that club cells, which are protective secretory cells in the epithelium of terminal and respiratory bronchioles, have regenerated after their initial ablation with naphthalene. Moreover, TGFluo and FLIM showed that LSCs preferentially engraft in injured bronchioles (compared to alveoli and to bronchioles of uninjured mice) where they participate to club cells regeneration (Figure 6.1b). This proof of principle experiment [18] showed that FND‐labeling of stem cells, combined to TGFluo and FLIM, allows in‐depth investigation of the optimal conditions of engraftment and regeneration in the targeted tissue. The method was recently extended to monitor the homing of human mesenchymal stem cells (MSCs) in miniature pigs [19], the respiratory system of this animal having human‐like immune responses. MSCs are self‐renewing, multipotent progenitor cells with the capacity to differentiate into distinct mesenchymal lineages. In humans, MSCs are mainly found in bone marrow, adipose, and placenta tissues. Owing to their multilineage differentiation potential, MSCs are considered as the most promising stem cells for therapy and regenerative medicine, which is a strong motivation to develop reliable methods to measure their biodistribution and pharmacokinetics in vivo in preclinical evaluations. Indeed, such methods will be crucial to determine what extent the transplanted MSCs home to the target organs, optimize the treatment and avoid inappropriate differentiation that can lead to cancer in the case of incorrect location. As in the previous study on mice [18], cell viability, immunomodulation and proliferation assays were conducted in vitro, and FND uptake by MSCs did not impact any of these properties. Then, for their investigation of the fate of FND‐labeled MSCs injected in miniature pigs, Su et al. [19] have taken advantage of the unique magnetic field sensitivity of NV‐center fluorescence [14], so that under a modulation of an external magnetic field, only the fluorescence intensity of FND varies and not the one of tissue autofluorescence. Therefore, the FND signal can be retrieved within a strong autofluorescent background by simple demodulation [15]. Using this magnetically modulated fluorescence (MMF) background‐free technique, in combination with TGFluo imaging, Su et al. [19] were able to quantify precisely the amount of FND‐containing human placenta choriodecidual membrane‐derived MSCs (pcMSCs) in each of the five main organs of miniature pigs. The animals were sacrificed at specific time points after their intravenous injection of FND‐labeled pcMSCs. Their main organs were then extracted, digested with acids under high pressure, and FND fluorescence was precisely quantified in each of them, thanks to MMF technique. Up to 70% of the FND‐labeled pcMSCs were found to reside in the lungs at 24–48 hours after FND‐labeled pcMSCs injection. In contrast, using free FND led to only 25% of them ending in the lungs. The next most important fraction was found in the liver (≈2% in both FND‐labeled MSCs or free FND). This quantitative information could only be obtained, for the first time, thanks to the unique magneto‐optical properties of NV centers in FND. The combination of MMF and FLIM/TGFluo [18, 19] represents a promising alternative to radioisotope labeling commonly used in stem‐cell tracking applications, at the single cell levels [17]. Noteworthy, FND does not alter the differentiation of both mouse and human embryonal carcinoma stem cells into neurons in vitro, as induced by conventional protocols [20].

The perfect photostability, absence of toxicity, and high fluorescence brightness of FND are also key properties to track rare cells such as circulating tumor cells or cancer stem cells (CSCs). The latter belong to a subpopulation of tumor cells that are resistant to current chemotherapy and radiotherapy treatments. Despite growing evidence of CSCs implication in tumor growth and recurrence, their isolation and eradication are still a challenge [21]. Conventional fluorescent markers are not chemically and photophysically stable enough to track CSCs on the few‐days (up to one month) period required for diagnosis and prognosis applications. Lin et al. [22] have shown that FND is a promising label for such a goal. They have used a human breast cancer cell line established from a patient and selected for their great capacity to form mammospheres (clumps of mammary gland cells), an indication for the presence of a stem cell population. Lin et al. [22] have discovered in in vitro assays that there are two populations of cells having spontaneously uptaken FND, a population FND⁺ with high content of FND, and another one FND⁻ with a low content. Incidentally, the fluorescence signal from FND⁻ cells, as measured by flow cytometry, decreased faster than the one of FND⁺ . This indicates that FND⁺ may belong to a slowly proliferating, quiescent cell subpopulation, possibly corresponding to a CSC phenotype. FND labeling was also compared to current CSC fluorescence labeling methods, and was found to outperform them in terms of (i) absence of genotoxicity and (ii) longer term tracking capability owing to a longer retention in cells [22]. This proof of principle experiment demonstrated the potential use of FND to track and find quiescent human CSCs.

We have mentioned studies showing that cells labeled with cytoplasmic FND can be retrieved in an organism on a long‐term scale, even at the single cell level and in a highly autofluorescent environment. While such experiments are carried out on sacrificed animals, detection of such FND‐labeled cells in a living organism is more challenging. However, Hui et al. [23] have extended the raster scanning TGFluo detection of FND to dynamical wide‐field imaging. To this aim, they used an intensified charge‐coupled device array detector and triggered its gate opening with some delay relative to the excitation lase pulse. This setup allowed them to track FND‐labeled mouse lung cancer cell motion at video rate in the blood stream of a living mouse after injection in its vein (Figure 6.2). Fluorescence imaging was then carried out near one of the main blood vessels of the mouse's ear. The autofluorescence (from sebaceous glands) was still present in TGFluo images, but it can be readily removed by post‐processed background subtraction so that the motion of FND‐labeled cells can be detected with a large signal‐to‐background ratio, which facilitates the trajectory extraction (Figure 6.2c). One potential application of TGFluo wide‐field microscopy of FND could be single particle tracking in a complex environment such as the brain. In Section 6.4 of this chapter, we present an alternative carbon‐based nanomaterial, i.e. single‐wall carbon nanotube, that is tracked individually thanks to its emission in the near‐infrared tissue transparency window, in order to probe the extracellular nanoscale environment in an intact brain tissue.

In this section, we have shown that FND, thanks to (i) its remarkable photophysical and magneto‐optical properties that allow background‐free imaging, and to (ii) its very low cellular toxicity, is a very promising label to optimize stem cell therapy protocols in preclinical research. Several studies have shown that FND can provide quantitative data that are hardly accessible by other methods. Furthermore, FND‐labeled cells can also be tracked individually in real time with background‐free wide‐field video‐microscopy, offering appealing prospects to identify rare cells like circulating tumor cells, directly in blood circulation.

6.3 Monitoring Inter and Intra Cellular Dynamics with Fluorescent Nanodiamonds

In the previous section, we have discussed several studies that filled up cells with FND and tracked their fate in organism like mice or pigs. However, FND can also be tracked at the single particle level in semi‐transparent organisms such as zebrafish [24], C‐elegans [25], or drosophila embryos [26]. The main motivation was that bright near‐infrared emitting FND reporters might help to understand the inner dynamics of a developing organism. For example, in Chang et al. [24], 100 nm‐size FND, coated with bovine serum albumin (BSA) to prevent aggregation, were microinjected into the yolk of a zebrafish embryo at the one‐cell stage. They found that single FND underwent unidirectional and stop‐and‐go traffic in the yolk cell with an average velocity of 0.3 μm s⁻¹ . Moreover, when incorporated into dividing cells, these particles could migrate into the fish's body as the embryos developed from larvae into adult fish. Interestingly, fishes injected with FND did not present abnormalities at the adult stage, meaning that FND did not interfere with embryogenesis.

The first attempt to track single FND in a whole organism was made by Igarashi et al. [27]. They developed a selective imaging method based on ODMR (cf. Section 6.1 for definition) to improve the image contrast of FND in vivo. Briefly, two wide‐field fluorescence images were recorded with or without a microwave (MW) modulation at the ODMR resonant frequency of 2.87 GHz. Subtracting two images (with MW on and MW off) pixel‐by‐pixel allowed them to remove the autofluorescence that is insensitive to MW and provides images displaying FND alone. Then, they performed long‐term tracking of single FND in both C. elegans and mice. In a following paper, they improved even further the temporal resolution of their apparatus (sampling rate increased up to 20 kHz) by using a spectrometer equipped with an avalanche photodiode [28]. As a proof‐of‐principle, they simply injected FND into the intestine of C. elegans and performed real‐time recording of FND without background fluorescence. Although the experiments reported in these articles were not addressing specific biological challenges, this selective imaging technique could be implemented for other living organisms and bring unexpected findings.

As previously discussed in Section 6.1, time‐gated imaging is an alternative approach to achieve background‐free detection of FND in whole organisms. For example, Kuo et al. [25] used it to investigate the intercellular transport of yolk lipoproteins in C. elegans (Figure 6.3a). FND were first noncovalently coated with green fluorescent protein (GFP)‐tagged yolk lipoprotein complexes (YLC) to for the GFP::YLC‐FND platform. Functionalized FNDs were then microinjected into the anterior intestinal cells near the pharynx. Worms were examined individually to identify the location of GFP::YLC‐FND in the specific cells or organs over 55 minutes after injection (Figure 6.3a). Results showed that the functionalized FND appeared in the posterior intestine immediately after administration. Then, they migrated into the pseudo‐coelom and the loop region of the gonad at 5 and 12 minutes postinjection, respectively. At 20 minutes postinjection, they became visible in the oocytes. Interestingly, an FND‐containing embryo was found at 55 minutes postinjection, indicating that oocytes can be fertilized and can normally develop into embryos.

Another example of single particle tracking of FND inside living organisms is the study of the embryogenesis in the drosophila [26]. In this article, the author introduced BSA‐coated FNDs into the embryo and investigated the FND diffusion in both furrow periplasm and subnuclear periplasms by SPT (Figure 6.3b). They observed that during cellularization at the posterior end of the drosophila embryos, the internalized FND in the blastoderm cells displayed two regimes of motion: free diffusion and molecular motor‐driven motion. By tracking the FND and extracting the trajectories, the authors determined a mean diffusion coefficient of 0.006 μm²  s⁻¹ in the furrow and 0.06 μm²  s⁻¹ in the sub‐nuclear periplasm. Although the velocity in these two compartments were similar (0.13 and 0.27 μm s⁻¹ respectively), the mean diffusion coefficient differs by one order of magnitude. The author therefore concluded that cytoskeletal networks in the furrow periplasm in more compact that in the subnuclear periplasm.

Although we previously described the use of FND tracking in fixed [18] or living organisms [25–27], the seminal article that introduced FND in biology a decade ago actually showed that this fluorescent particle can be used to study intracellular dynamics [29, 30]. This approach requires to internalize single FND inside a cell and track them over time using a diffraction‐limited optical system. The first proof of principle was done in cancer cell line, which are easy to grow and maintain. Moreover, due to their intrinsic proliferative behavior, exogeneous compounds are easy to internalize by endocytosis. In a subsequent article, the same group showed that bright 35 nm‐sized FND can be tracked in 3D in a reproducible manner [12]. Figure 6.4a displays a typical result of a bare FND moving inside a HeLa cell.

Another application of SPT of FND inside living cells is described in Liu et al. [31]. In this work, FND were conjugated to the transforming growth factor (TGF) in order to target the TGF‐β membrane receptors and track their motion in 3D. The authors showed that the TGF‐β receptor presents three different regimes of motion: immobile, intermediate, and fast diffusion (Figure 6.4b). After treatment with small molecule kinase inhibitors (SMI), the fraction of the immobile population significantly decreased compared to the one of intermediate and fast populations. This result is consistent with the fact that SMI releases the TGF‐β receptor from a larger binding complex, therefore increasing its diffusion on the membrane. The author claimed this data proves that immobilized TGF‐β is essential for active signaling. Overall, this work shows that FND can be used as specific tagging nanotool of endogenous proteins in living cells for the study of transmembrane signaling dynamics.

Interestingly, by recording the ODMR spectra over time, McGuinness et al. [32] were able to track the orientation of a single FND entrapped in a living cell. The author applied a uniform magnetic field to live HeLa cells containing 50‐nm FND, each of which hosts a single NV⁻ center. By closely analyzing the resonance frequency shift in the ODMR spectra, information on the rotational motions of single FND in the cell was inferred. Thanks to the perfect photostability of FND, the authors were able to continuously track the orientation change over 16 hours.

Until recently, single particle tracking of FND has only been conducted in cancer cell lines. Although, this biological system can be useful for drug discovery or fundamental studies on intracellular dynamics, it falls short to address some other questions like brain disease‐related ones requiring culture of neurons dissociated from mouse embryos. A recent study proved that SPT of FND can be easily conducted in neuron culture with a high enough sensitivity to detect intraneuronal transport abnormalities [1]. Previous attempts to quantify intraneuronal transport have been using fluorescent proteins to label organelles [33]. However, this approach faces several limitations: low transfection yield, uncontrolled protein expression, photobleaching, and cytotoxicity. Fluorescent semiconductor nanocrystals (quantum dots, QDs) is another type of fluorescent reporter that was used to monitor various cellular events [34], but attempts to use QDs to study intraneuronal transport have been limited to the specific model of long axons of dorsal root neurons cultured in microfluidic devices [35]. Moreover, QDs blinking impedes high spatiotemporal resolution tracking and therefore biases transport parameter measurements [1]. Considering the intrinsic limitations of other potentially useful reporters, FND unique properties of high brightness, photostability, and absence of cytotoxicity, make them a tool of choice to detect abnormalities of intraneuronal transport.

Haziza et al. [1] has developed a novel quantitative assay based on tracking of single 30 nm‐sized FND in mouse hippocampal neurons (Figure 6.5a). They took advantage of the endocytosis mechanism to introduce FND into neuronal branches and recorded FND trajectories with Total Internal Reflection Fluorescence (TIRF) video microscopy at 50 ms temporal resolution, giving a spatial localization accuracy of 12 nm. They were then able to reconstruct the entire trajectories with a tracking software (Figure 6.5b). Using a custom‐made algorithm, they extracted relevant transport readouts: velocity (in μm/s), run length (in μm), processivity (in seconds), pausing time (in seconds), pausing frequency (events per minute) and the diffusion coefficient (in μm² /s). Interestingly, they proved the superiority of FND over QD to reliably report transport parameters and therefore emphasizes that the perfect photostability of FND is key for this approach [1]. To prove the sensitivity of the FND‐tracking assay, the author designed three experiments with increasing complexity. They first used a pharmacological drug that impact the microtubule‐based intraneuronal transport, but at nanomolar concentrations. Then, as a validation of the ability of their technique to record abnormal intraneuronal transport, they incubated neuron culture with sub‐micromolar concentrations of amyloid‐β (Aβ) peptide, a well‐known molecular player Alzheimer's in disease [36, 37] (Figure 6.5c). This experiment suggests that the FND‐tracking assay could be used to screen drugs capable of rescuing Alzheimer's disease phenotype. Eventually, as a proof‐of‐principle, they applied their method on two transgenic mouse lines that mimic small variation (≈+30%) in protein concentration found in brains of patients [1]. In both cases, the FND‐tracking assay was sensitive enough to detect small modifications of the intraneuronal transport parameters in the transgenic neurons.

In this section, we have discussed the use of FND at the single particle level. Single FND tracking can be used in the whole organism like C‐elegans or drosophila embryos and led to important findings on the developmental organization. In addition, SPT of FND can be performed at the single cell levels. Historically done in cancer cell lines for basic research, the unique photophysical properties of FND and their biocompatibility brought them into the field of neurosciences and brain diseases. A novel FND‐tracking assay that monitor the endosomal transport inside neuronal branches was recently reported. It was proved to be sensitive enough to detect subtle change in gene expression, as the ones found in the brain of patient with autism or Alzheimer's disease. For the first time, a nanoparticle‐based assay is able to directly measure a functional impact of genetic risk factors found in patient with brain disease, opening the door for applications in translational nanomedicine.

6.4 Single‐Walled Carbon Nanotubes: A Near‐Infrared Optical Probe of the Nanoscale Extracellular Space in Live Brain Tissue

Some of the current challenges for a better understanding of fundamental processes in biology and for applications in medical science concern the development of imaging nanoprobes and chemical nanosensors which are not limited to single cell studies, but applicable within live intact tissues. Nanoprobes and nanosensors must ideally display intense and specific signals reporting (related to) local physical and/or chemical environments. The elaboration of new drug delivery strategies shares similar requirements since drug transporters must be able to carry multiple functions inside living organisms including (i) the recognition of a specific cellular/molecular target; (ii) the actuation of the drug itself and if needed; (iii) the signalization of their tissue localization. Whether we are dealing with drug transporters, imaging nanoprobes, or chemical nanosensors, all of them thus need to display excellent accessibility within intact tissues.

For these applications, the development of spherical particles as multifunctional objects encounters contradictory requirements: they should be small enough for tissue accessibility and large enough for carrying the multiple functions mentioned above, i.e. targeting capabilities and/or drug delivery, chemical sensing signals and imaging contrast through tissues. Therefore, the combination of optical, chemical, and penetration capabilities in tissue are usually not fulfilled together with spherical nanoparticles. Because they display extreme length‐to‐diameter aspect ratios as well as unique optical/chemical/mechanical properties, single‐walled carbon nanotubes (SWCNT) constitute a promising platform to fulfill these requirements.

SWCNT distinguish themselves from double‐walled carbon nanotubes and multi‐walled carbon nanotubes by their single‐layer cylindrical sidewall structure, which provides them more finely tuned physical and chemical properties for applications. They are characterized by the thinnest diameters (≈1–3 nm) and have lengths of hundreds of nanometers up to few micrometers, conferring them unusual aspect ratios. SWCNT display strong optical resonances and the discovery of their near‐infrared (NIR) luminescence [38, 39] from semiconducting species paved the way to their detection at the single nanotube level in liquid environments [40–42]. Individual semiconducting SWCNT exhibit luminescence with large Stokes shifts in the NIR window (typically c. 900–1400 nm), which falls in the therapeutic window of biological tissues (typically 650–1400 nm) and is thus potentially free of autofluorescence coming from biological structures. SWCNT optical properties, which are based on excitonic processes, are now rather well understood [43–45]. Despite their low luminescence quantum efficiencies [46–48], water‐solubilized SWCNT display exceptionally high luminescence rates for a single nano‐object [41] due to remarkably large absorption cross‐sections [49–53]. They also display exceptional luminescence signal stabilities (tens of min [54]) with negligible blinking and photobleaching. Using suitable encapsulation strategies, fluorescence emission of SWCNT can be highly stable, allowing long‐term imaging in aqueous environments at the single nanotube level [42, 54], but also intracellular tracking of single nanotube in cultured cells [55–57]. Interestingly, depending on their encapsulating agents, SWCNT photoluminescence can be extremely sensitive to their chemical environment [42, 58, 59], which opens the way to sensing capabilities in biological environments [60–64]. Moreover, their brightness, photostability, and spectral imaging range [38, 65–67] make SWCNT unique probes at the ensemble level for imaging the whole animal vascular systems [68, 69].

In the following, we will describe how the unique combination of SWCNT diffusion and optical properties within complex environments can be exploited at the single nanotube level in order to access live intact tissue structures and probe their nanoscale environment. Diffusion and optical imaging being inherent to the aforementioned applications, it will be possible to extend the following concepts to other developments such as chemical nanosensors, optical contrast agents, or drug delivery systems within living biological tissues.

Molecular diffusion processes are key for the function of living matter [70, 71]. However, they occur within complex environments characterized by heterogeneous nanoscale organizations and chemical contents, which make it difficult to dissect. In this context, single‐molecule optical microscopy has been crucial over the last decades to study the spatio‐temporal heterogeneity of the biological complexity. It provides the ability to probe the full population of diffusive behaviors, including the presence of minority populations (e.g. localized, anomalous, or Brownian diffusion), in multiple environments spanning from nanometer‐ to micrometer‐scale resolution. A widely used approach to studying molecular diffusion is the super‐localization technique, based on the determination of a diffraction‐limited emitter position with nanometer precision [72, 73] Because biological constituents are generally neither detectable nor resolvable at the molecular scales, single‐molecule microscopy commonly uses highly luminescent nano‐emitters that probe their biological structures while being efficiently detected by an optical microscope. This is usually performed in isolated cells or very thin tissue preparations (up to a few micrometers) at visible or eventually far‐red wavelengths using organic dyes, autofluorescent proteins or eventually small emitting nanoparticles. For applications in thick intact biological tissues, scattering and absorption of visible light complicates single molecule detection such that single‐molecule imaging within samples thicker than isolated cells is only emerging and is mostly limited to a few cellular layers [74, 75]. To access the optical window of biological tissues, i.e. where scattering and absorption of tissue are minimal, the use of nanoprobes with optical response in the NIR is thus crucial, and SWCNT match this requirement.

Prior to their use as biological nanoprobes, SWCNT have to be biocompatible. In liquid environment, the photoluminescence of SWCNT is obtained by coating their surface with surfactants or polymers. Although surfactants like sodium dodecylbenzene sulfonate or bile salts are known to provide the best luminescing SWCNT in aqueous environments [76, 77], their use for biological applications should be avoided as they alter the integrity of cellular membranes. Several biocompatible coatings were investigated, and among them, phospholipid‐polyethylene glycol (PL‐PEG) provided high luminescence, low toxicity [68], as well as minimal nonspecific interaction with cellular membrane [78]. The use of PL‐PEG allows long‐term imaging directly within live tissue [2] and long circulation time in the vascular system in vivo [69]. For applications where interactions with cellular constituent are sought, in particular for sensing applications, the use of short DNA sequences [60, 62], biomolecules [61], or biocompatible polymers [63, 64, 79] have provided interesting biocompatible routes for suspending luminescent SWCNT with sensing capabilities.

Several wavelengths that fall in the biological “transparency” window can be used to excite SWCNT in tissues. Considering the widely used (6, 5) SWCNT chirality,¹Danné et al. [80] investigated the possibility of exciting them in brain tissues at the single nanotube level by continuous‐wave (cw) lasers at their second‐order excitonic transition (568 nm), their K‐momentum exciton‐phonon sideband (845 nm), or through up‐conversion (1064 nm). The effects of tissue scattering, absorption, autofluorescence, and temperature increase induced by excitation light were systematically examined. Video‐rate imaging of single (6, 5) SWCNT diffusing in liquid environments upon cw up‐conversion excitation at 1064 nm was first demonstrated and the comparison between different excitation wavelength to obtain identical photoluminescence rates from single biocompatible PL‐PEG coated (6, 5) SWCNT was performed. This allowed a realistic determination of the best excitation strategy to detect single (6, 5) SWCNT in live brain tissue. Danné et al. [80] found that despite its most red‐shifted wavelength, care should be taken when up‐conversion aims at being used to image single SWCNT at video‐rate in live tissues because of the temperature rise induced by laser absorption of the tissue. They concluded that excitation at the K‐momentum exciton‐phonon sideband (845 nm) is currently the optimal strategy for imaging single (6, 5) SWCNT while further fundamental understanding of the processes underlying up‐conversion of SWCNT might help optimizing excitation efficiencies of single SWCNT.

To serve as biological nanoprobes in tissues, SWCNT long 1D shape inevitably raised concerns about tissue penetration. A first insight about their potential for tissue penetration was provided through the discovery that long SWCNT can move by reptation, or “snake‐like motion,” in crowded environments [54] (Figure 6.6). The extent of SWCNT diffusion in such environments is indeed a function of their length, diameter, and of environment crowdedness [2]. More precisely, the persistence length (L _p) or rigidity of SWCNT depends on their diameter and was previously characterized using thermal bending statistics [81]. It ranges from a few tens of μm up to several hundreds of μm. It follows that when SWCNT are immersed in crowded environments, they undergo Brownian or random thermal motion according to two distinct diffusive regimes, depending on their length, persistence length, and the network characteristic dimension (ξ) [54]. A first regime, proposed by Odijk [82], takes place when the SWCNT length is larger than the independent deflection length λ = (L _p ξ² )^1/3 , in which case SWCNT will undergo reptation as introduced by de Gennes [83]. A second regime, predicted by Doi [84], happens when SWCNT length is smaller than the independent deflection length λ. For Doi regime, in contrast to Odijk regime, the diffusion is completely independent of the flexibility and can be modeled assuming fully rigid SWCNT. The transition between these two regimes were observed experimentally by Fakhri et al. [54] by analyzing the diffusion of individual SWCNT of different rigidities (i.e. different diameters) and lengths in aqueous gels. These types of behaviors later lead to the observation obtained at the ensemble level by Jena et al. [85], that SWCNT unusual aspect ratios contribute to enhancing their penetration within multicellular tumor spheroids, but this enhancement was also found dependent on the encapsulation strategy [85, 86]. Godin et al. [2] pushed these concepts a step further by performing single nanotubes studies in live brain tissues. They showed that the analysis of the movement of single PL‐PEG coated SWCNT can be used to unravel the extracellular dimensions and fluidity of live brain tissues. They indeed demonstrated that upon injection into the lateral cerebroventricles of young rat brains, PL‐PEG coated SWCNT diameters were small enough to enabling diffusion into small brain compartments, but at the same time, that the SWCNT length is sufficiently long to slow down SWCNT movements and allow them to extensively explore their submicroscopic environment. In addition, due of their photostability and NIR emission properties, single SWCNT could be imaged in the extracellular space of acute brain slices in different areas of the sagittal brain, at imaging depths of a few tens of μm up to ~100 μm. Their movements could be tracked for long periods of time at video rate, allowing recording trajectories that are orders of magnitude longer than trajectories obtained by tracking small isotropic probes (e.g. dye molecules, antibodies, latex beads, and quantum dots). Super localization of the nanotube positions provided the first super‐resolved maps of the extracellular space of live brain tissues (down to 40 nm resolution, Figure 6.7), demonstrating that the extracellular space is larger than previously thought in live brain, as recently suggested by electron microscopy in cryo‐fixed tissues [87]. In addition, single SWCNT tracking in live tissues revealed the heterogeneity of the extracellular space local viscosity and its dependence on extracellular matrix components.

Altogether, these findings indicate that SWCNT constitutes a new platform to envision a better understanding of the extracellular space of live tissues, through the combination of unique tissue accessibility and optical properties for deep‐tissue nanoimaging, but surely in future for local chemical sensing capabilities in local tissue areas. These new nanoreporters are based on SWCNT and will confidently disseminate in biology, grounding new fundamental concepts in organ development, physiology and pathophysiology. We can also envision that such concepts can be applied for the development of novel drug delivery strategies based on 1D nanoscale transporters.

6.5 Conclusion

In this chapter, we have highlighted recent works that applied photoluminescent carbon nanomaterials, mainly FND and single‐walled carbon nanotubes (SWCNT), to dissect cellular and organism dynamics. The unique combination of SWCNT diffusion and optical properties can be exploited at the single nanotube level to probe nanoscale environment in living tissues, such as a mouse brain. Although one can argue that FND might suffer from its too‐large size (typically 30–100 nm), several groups have made progress to reduce it down to 10 nm without significantly compromising their photophysical properties [88, 89]. In fact, researchers are benefiting from the perfect photostability, the unique magneto‐optical property, the easy surface chemistry activation, and the absence of toxicity of FND and thoroughly used them at different biological scales. FND can help tracking cells in organisms like mouse or miniature pigs [18, 19], or can be used to monitor membrane receptor dynamics [31] and intraneuronal transport [1]. They may also serve as gene and drug delivery vehicles by modifying their surface chemistry [90, 91]. Altogether, despite many challenges, carbon nanomaterials have proven themselves as highly promising imaging tools to screen cellular and organism dynamics. The next step would be to translate these methodologies and findings from labs to clinical applications such as diagnosis and therapy.

References

1. 1 Haziza, S., Mohan, N., Loe‐Mie, Y. et al. (2017). Fluorescent nanodiamond tracking reveals intraneuronal transport abnormalities induced by brain‐disease‐related genetic risk factors. Nat. Nanotechnol. 12 (4): 322–328.
2. 2 Godin, A.G., Varela, J.A., Gao, Z. et al. (2016). Single‐nanotube tracking reveals the nanoscale organization of the extracellular space in the live brain. Nat. Nanotechnol. 12 (3): 238–243.
3. 3 Tan, L., Wu, T., Tang, Z.‐W. et al. (2016). Water‐soluble photoluminescent fullerene capped mesoporous silica for pH‐responsive drug delivery and bioimaging. Nanotechnology 27 (31): 315104.
4. 4 Zhang, H., Zhang, H., Aldalbahi, A. et al. (2017). Fluorescent biosensors enabled by graphene and graphene oxide. Biosens. Bioelectron. 89: 96–106.
5. 5 Wang, J. and Qiu, J. (2016). A review of carbon dots in biological applications. J. Mater. Sci. 51 (10): 4728–4738.
6. 6 Faklaris, O., Garrot, D., Joshi, V. et al. (2008). Detection of single Photoluminescent diamond nanoparticles in cells and study of the internalization pathway. Small 4 (12): 2236–2239.
7. 7 Faklaris, O., Joshi, V., Irinopoulou, T. et al. (2009). Photoluminescent diamond nanoparticles for cell Labeling: study of the uptake mechanism in mammalian cells. ACS Nano 3 (12): 3955–3962.
8. 8 Fang, C.‐Y., Vaijayanthimala, V., Cheng, C.‐A. et al. (2011). The exocytosis of fluorescent Nanodiamond and its use as a long‐term cell tracker. Small 7 (23): 3363–3370.
9. 9 Huang, Y.‐A., Kao, C.‐W., Liu, K.‐K. et al. (2014). The effect of fluorescent nanodiamonds on neuronal survival and morphogenesis. Sci. Rep. 4: 6919.
10. 10 Simpson, D.A., Morrisroe, E., McCoey, J.M. et al. (2017). Non‐neurotoxic Nanodiamond probes for Intraneuronal temperature mapping. ACS Nano 11 (12): 12077–12086.
11. 11 Paget, V., Sergent, J.A., Grall, R. et al. (2013). Carboxylated nanodiamonds are neither cytotoxic nor genotoxic on liver, kidney, intestine and lung human cell lines. Nanotoxicology 5390: 1–11.
12. 12 Chang, Y.‐R., Lee, H.‐Y., Chen, K. et al. (2008). Mass production and dynamic imaging of fluorescent nanodiamonds. Nat. Nanotechnol. 3 (5): 284–288.
13. 13 Dantelle, G., Slablab, A., Rondin, L. et al. (2010). Efficient production of NV colour centres in nanodiamonds using high‐energy electron irradiation. J. Lumin. 130 (9): 1655–1658.
14. 14 Doherty, M.W., Manson, N.B., Delaney, P. et al. (2013). The nitrogen‐vacancy colour centre in diamond. Phys. Rep. 528 (1): 1–45.
15. 15 Sarkar, S.K., Bumb, A., Wu, X. et al. (2014). Wide‐field in vivo background free imaging by selective magnetic modulation of nanodiamond fluorescence. Biomed. Opt. Express 5 (4): 1190–1202.
16. 16 Greffet, J.‐J., Hugonin, J.‐P., Besbes, M. et al. (2011). Diamond particles as nanoantennas for nitrogen‐vacancy color centers. http://arXiv.org. Quantum Phys. 18: 1–4.
17. 17 Nguyen, P.K., Riegler, J., and Wu, J.C. (2014). Stem cell imaging: from bench to bedside. Cell Stem Cell 14 (4): 431–444.
18. 18 Wu, T., Tzeng, Y.‐K., Chang, W.‐W. et al. (2013). Tracking the engraftment and regenerative capabilities of transplanted lung stem cells using fluorescent nanodiamonds. Nat. Nanotechnol. 8 (9): 682–689.
19. 19 Su, L.‐J., Wu, M.‐S., Hui, Y.Y. et al. (2017). Fluorescent nanodiamonds enable quantitative tracking of human mesenchymal stem cells in miniature pigs. Sci. Rep. 7 (February): 45607.
20. 20 Hsu, T.‐C., Liu, K.‐K., Chang, H.‐C. et al. (2015). Labeling of neuronal differentiation and neuron cells with biocompatible fluorescent nanodiamonds. Sci. Rep. 4 (1): 5004.
21. 21 Batlle, E. and Clevers, H. (2017). Cancer stem cells revisited. Nat. Med. 23 (10): 1124–1134.
22. 22 Lin, H.‐H., Lee, H.‐W., Lin, R.‐J. et al. (2015). Tracking and finding slow‐proliferating/quiescent cancer stem cells with fluorescent Nanodiamonds. Small 11 (34): 4394–4402.
23. 23 Hui, Y.Y., Su, L.‐J., Chen, O.Y. et al. (2014). Wide‐field imaging and flow cytometric analysis of cancer cells in blood by fluorescent nanodiamond labeling and time gating. Sci. Rep. 4 (1): 5574.
24. 24 Chang, C.‐C., Zhang, B., Li, C.‐Y., et al. (2012). Exploring cytoplasmic dynamics in zebrafish yolk cells by single particle tracking of fluorescent nanodiamonds. Adv Photonics Quantum Comput Mem Commun V. 8272:SPIE‐SPIE.
25. 25 Kuo, Y., Hsu, T.‐Y., Wu, Y.‐C., and Chang, H.‐C. (2013). Fluorescent nanodiamond as a probe for the intercellular transport of proteins in vivo. Biomaterials 34 (33): 8352–8360.
26. 26 Simpson, D.A., Thompson, A.J., Kowarsky, M. et al. (2014). In vivo imaging and tracking of individual nanodiamonds in drosophila melanogaster embryos. Biomed. Opt. Express 5 (4): 1250.
27. 27 Igarashi, R., Yoshinari, Y., Yokota, H. et al. (2012). Real‐time background‐free selective imaging of fluorescent nanodiamonds in vivo. Nano Lett. 12 (11): 5726–5732.
28. 28 Yoshinari, Y., Mori, S., Igarashi, R. et al. (2015). Optically detected magnetic resonance of Nanodiamonds in vivo; implementation of selective imaging and fast sampling. J. Nanosci. Nanotechnol. 15 (2): 1014–1021.
29. 29 Fu, C.‐C., Lee, H.‐Y., Chen, K. et al. (2007). Characterization and application of single fluorescent nanodiamonds as cellular biomarkers. Proc. Natl. Acad. Sci. 104 (3): 727–732.
30. 30 Hui, Y.Y., Hsiao, W.W.‐W., Haziza, S. et al. (2017). Single particle tracking of fluorescent nanodiamonds in cells and organisms. Curr. Opin. Solid State Mater. Sci. 21 (1): 35–42.
31. 31 Liu, W., Yu, F., Yang, J. et al. (2016). 3D single‐molecule imaging of Transmembrane Signaling by targeting Nanodiamonds. Adv. Funct. Mater. 26 (3): 365–375.
32. 32 McGuinness, L.P., Yan, Y., Stacey, A. et al. (2011). Quantum measurement and orientation tracking of fluorescent nanodiamonds inside living cells. Nat. Nanotechnol. 6 (6): 358–363.
33. 33 Kwinter, D.M., Lo, K., Mafi, P., and Silverman, M.A. (2009). Dynactin regulates bidirectional transport of dense‐core vesicles in the axon and dendrites of cultured hippocampal neurons. Neuroscience 162 (4): 1001–1010.
34. 34 Pinaud, F., Clarke, S., Sittner, A., and Dahan, M. (2010). Probing cellular events, one quantum dot at a time. Nat. Methods 7 (4): 275–285.
35. 35 Mudrakola, H.V., Zhang, K., and Cui, B. (2009). Optically resolving individual microtubules in live axons. Structure 17 (11): 1433–1441.
36. 36 Stokin, G.B., Lillo, C., Falzone, T.L. et al. (2005). Axonopathy and transport deficits early in the pathogenesis of Alzheimer's disease. Science 307 (5713): 1282–1288.
37. 37 Encalada, S.E. and Goldstein, L.S.B. (2014). Biophysical challenges to axonal transport: motor‐cargo deficiencies and Neurodegeneration. Annu. Rev. Biophys. 43 (1): 141–169.
38. 38 O'Connell, M.J., Bachilo, S.M., Huffman, C.B. et al. (2002). Band gap fluorescence from individual single‐walled carbon nanotubes. Science 297 (5581): 593–596.
39. 39 Bachilo, S.M., Strano, M.S., Kittrell, C. et al. (2002). Structure‐assigned optical spectra of single‐walled carbon nanotubes. Science 298 (5602): 2361–2366.
40. 40 Tsyboulski, D.A., Bachilo, S.M., and Weisman, R.B. (2005). Versatile visualization of individual single‐walled carbon nanotubes with near‐infrared fluorescence microscopy. Nano Lett. 5 (5): 975–979.
41. 41 Tsyboulski, D.A., Rocha, J.D.R., Bachilo, S.M. et al. (2007). Structure‐dependent fluorescence efficiencies of individual single‐walled carbon nanotubes. Nano Lett. 7 (10): 3080–3085.
42. 42 Cognet, L., Tsyboulski, D.A., Rocha, J.‐D.R. et al. (2007). Stepwise quenching of Exciton fluorescence in carbon nanotubes by single‐molecule reactions. Science 316 (5830): 1465–1468.
43. 43 Wang, F. (2005). The optical resonances in carbon nanotubes Arise from Excitons. Science 308 (5723): 838–841.
44. 44 Maultzsch, J., Pomraenke, R., Reich, S. et al. (2005). Exciton binding energies in carbon nanotubes from two‐photon photoluminescence. Phys. Rev. B 72 (24): 241402.
45. 45 Perebeinos, V. and Phonon, A.P. (2008). Electronic Nonradiative decay mechanisms of Excitons in carbon nanotubes. Phys. Rev. Lett. 101 (5): 57401.
46. 46 Lefebvre, J., Austing, D.G., Bond, J., and Finnie, P. (2006). Photoluminescence imaging of suspended single‐walled carbon nanotubes. Nano Lett. 6 (8): 1603–1608.
47. 47 Berciaud, S., Cognet, L., and Lounis, B. (2008). Luminescence decay and the absorption cross section of individual single‐walled carbon nanotubes. Phys. Rev. Lett. 101 (7): 77402.
48. 48 Crochet, J., Clemens, M., and Hertel, T. (2007). Quantum yield heterogeneities of aqueous single‐wall carbon nanotube suspensions. J. Am. Chem. Soc. 129 (26): 8058–8059.
49. 49 Schöppler, F., Mann, C., Hain, T.C. et al. (2011). Molar extinction coefficient of Single‐Wall carbon nanotubes. J. Phys. Chem. C 115 (30): 14682–14686.
50. 50 Oudjedi, L., Parra‐Vasquez, A.N.G., Godin, A.G. et al. (2013). Metrological investigation of the (6,5) carbon nanotube absorption cross section. J. Phys. Chem. Lett. 4 (9): 1460–1464.
51. 51 Christofilos, D., Blancon, J.C., Arvanitidis, J. et al. (2012). Optical imaging and absolute absorption cross section measurement of individual nano‐objects on opaque substrates: single‐wall carbon nanotubes on silicon. J. Phys. Chem. Lett. 3 (9): 1176–1181.
52. 52 Vialla, F., Roquelet, C., Langlois, B. et al. (2013). Chirality dependence of the absorption cross section of carbon nanotubes. Phys. Rev. Lett. 111 (13): 137402.
53. 53 Streit, J.K., Bachilo, S.M., Ghosh, S. et al. (2014). Directly measured optical absorption cross sections for structure‐selected single‐walled carbon nanotubes. Nano Lett. 14 (3): 1530–1536.
54. 54 Fakhri, N., MacKintosh, F.C., Lounis, B. et al. (2010). Brownian motion of stiff filaments in a crowded environment. Science 330 (6012): 1804–1807.
55. 55 Cherukuri, P., Bachilo, S.M., Litovsky, S.H., and Weisman, R.B. (2004). Near‐infrared fluorescence microscopy of single‐walled carbon nanotubes in phagocytic cells. J. Am. Chem. Soc. 126 (48): 15638–15639.
56. 56 Reuel, N.F., Dupont, A., Thouvenin, O. et al. (2012). Three‐dimensional tracking of carbon nanotubes within living cells. ACS Nano 6 (6): 5420–5428.
57. 57 Fakhri, N., Wessel, A.D., Willms, C. et al. (2014). High‐resolution mapping of intracellular fluctuations using carbon nanotubes. Science 344 (6187): 1031–1035.
58. 58 Cambré, S., Santos, S.M., Wenseleers, W. et al. (2012). Luminescence properties of individual empty and water‐filled single‐walled carbon nanotubes. ACS Nano. 6 (3): 2649–2655.
59. 59 Duque, J.G., Oudjedi, L., Crochet, J.J. et al. (2013). Mechanism of electrolyte‐induced brightening in single‐wall carbon nanotubes. J. Am. Chem. Soc. 135 (9): 3379–3382.
60. 60 Heller, D.A., Jin, H., Martinez, B.M. et al. (2009). Multimodal optical sensing and analyte specificity using single‐walled carbon nanotubes. Nat. Nanotechnol. 4 (2): 114–120.
61. 61 Jin, H., Heller, D.A., Kalbacova, M. et al. (2010). Detection of single‐molecule H₂O₂ signalling from epidermal growth factor receptor using fluorescent single‐walled carbon nanotubes. Nat. Nanotechnol. 5 (4): 302–309.
62. 62 Heller, D.A., Pratt, G.W., Zhang, J. et al. (2011). Peptide secondary structure modulates single‐walled carbon nanotube fluorescence as a chaperone sensor for nitroaromatics. Proc. Natl. Acad. Sci. 108 (21): 8544–8549.
63. 63 Kruss, S., Landry, M.P., Vander Ende, E. et al. (2014). Neurotransmitter detection using corona phase molecular recognition on fluorescent single‐walled carbon nanotube sensors. J. Am. Chem. Soc. 136 (2): 713–724.
64. 64 Budhathoki‐Uprety, J., Langenbacher, R.E., and Jena, P.V. (2017). A carbon nanotube optical sensor reports nuclear entry via a noncanonical pathway. ACS Nano 11 (4): 3875–3882.
65. 65 Ju, S.Y., Kopcha, W.P., and Papadimitrakopoulos, F. (2009). Brightly fluorescent single‐walled carbon nanotubes via an oxygen‐excluding surfactant organization. Science 323 (5919): 1319–1323.
66. 66 Diao, S., Hong, G., Antaris, A.L. et al. (2015). Biological imaging without autofluorescence in the second near‐infrared region. Nano Res. 8 (9): 3027–3034.
67. 67 Hong, G., Diao, S., Antaris, A.L., and Dai, H. (2015). Carbon Nanomaterials for biological imaging and Nanomedicinal therapy. Chem. Rev. 115 (19): 10816–10906.
68. 68 Welsher, K., Liu, Z., Sherlock, S.P. et al. (2009). A route to brightly fluorescent carbon nanotubes for near‐infrared imaging in mice. Nat. Nanotechnol. 4 (11): 773–780.
69. 69 Hong, G., Diao, S., Chang, J. et al. (2014). Through‐skull fluorescence imaging of the brain in a new near‐infrared window. Nat. Photonics 8 (9): 723–730.
70. 70 Berg, H.C. (1993). Random Walks in Biology . Princeton University Press.
71. 71 Dusenbery, D.B. (2009). Living at Micro Scale: The Unexpected Physics of Being Small . Harvard University Press.
72. 72 Thompson, R.E., Larson, D.R., and Webb, W.W. (2002). Precise Nanometer localization analysis for individual fluorescent probes. Biophys. J. 82 (5): 2775–2783.
73. 73 Yildiz, A. (2003). Myosin V walks hand‐over‐hand: single Fluorophore imaging with 1.5‐nm localization. Science 300 (5628): 2061–2065.
74. 74 Biermann, B., Sokoll, S., Klueva, J. et al. (2014). Imaging of molecular surface dynamics in brain slices using single‐particle tracking. Nat. Commun. 5: 3024.
75. 75 Varela, J.A., Dupuis, J.P., Etchepare, L. et al. (2016). Targeting neurotransmitter receptors with nanoparticles in vivo allows single‐molecule tracking in acute brain slices. Nat. Commun. 7: 10947.
76. 76 Wenseleers, W., Vlasov, I.L., Goovaerts, E. et al. (2004). Efficient isolation and solubilization of pristine single‐walled nanotubes in bile salt micelles. Adv. Funct. Mater. 14 (11): 1105–1112.
77. 77 Duque, J.G., Pasquali, M., Cognet, L., and Lounis, B. (2009). Environmental and synthesis‐dependent luminescence properties of individual single‐walled carbon nanotubes. ACS Nano 3 (8): 2153–2156.
78. 78 Gao, Z., Danné, N., Godin, A. et al. (2017). Evaluation of different single‐walled carbon nanotube surface coatings for single‐particle tracking applications in biological environments. Nanomaterials 7 (11): 393.
79. 79 Bisker, G., Dong, J., Park, H.D. et al. (2016). Protein‐targeted corona phase molecular recognition. Nat. Commun. 7: 10241.
80. 80 Danné, N., Godin, A.G., Gao, Z. et al. (2018). Comparative analysis of photoluminescence and Upconversion emission from individual carbon nanotubes for bioimaging applications. ACS Photonics 5 (2): 359–364.
81. 81 Fakhri, N., Tsyboulski, D.A., Cognet, L. et al. (2009). Diameter‐dependent bending dynamics of single‐walled carbon nanotubes in liquids. Proc. Natl. Acad. Sci. 106 (34): 14219–14223.
82. 82 Odijk, T. (1983). On the statistics and dynamics of confined or entangled stiff polymers. Macromolecules 16 (8): 1340–1344.
83. 83 de Gennes, P.G. (1971). Reptation of a polymer chain in the presence of fixed obstacles. J. Chem. Phys. 55 (2): 572–579.
84. 84 Doi, M. (1975). Rotational relaxation time of rigid rod‐like macromolecule in concentrated solution. J. Phys. 36 (7–8): 607–611.
85. 85 Jena, P.V., Shamay, Y., Shah, J. et al. (2016). Photoluminescent carbon nanotubes interrogate the permeability of multicellular tumor spheroids. Carbon N. Y. 97: 99–109.
86. 86 Wang, Y., Bahng, J.H., Che, Q. et al. (2015). Anomalously fast diffusion of targeted carbon nanotubes in cellular spheroids. ACS Nano 9 (8): 8231–8238.
87. 87 Korogod, N., Petersen, C.C., and Knott, G.W. (2015). Ultrastructural analysis of adult mouse neocortex comparing aldehyde perfusion with cryo fixation. Elife 4: e05793.
88. 88 Mohan, N., Tzeng, Y.‐K., Yang, L. et al. (2010). Sub‐20‐nm fluorescent Nanodiamonds as Photostable biolabels and fluorescence resonance energy transfer donors. Adv. Mater. 22 (7): 843–847.
89. 89 Boudou, J.‐P., Tisler, J., Reuter, R. et al. (2013). Fluorescent nanodiamonds derived from HPHT with a size of less than 10 nm. Diamond Relat. Mater. 37: 80–86.
90. 90 Alhaddad, A., Adam, M.‐P., Botsoa, J. et al. (2011). Nanodiamond as a vector for siRNA delivery to Ewing sarcoma cells. Small 7 (21): 3087–3095.
91. 91 Slegerova, J., Hajek, M., Rehor, I. et al. (2015). Designing the nanobiointerface of fluorescent nanodiamonds: highly selective targeting of glioma cancer cells. Nanoscale 7 (2): 415–420.