Bioorthogonal chemical imaging of solid lipid nanoparticles with minimal labeling by stimulated Raman scattering microscopy

Abstract

Solid lipid nanoparticles (SLNs) are a state‐of‐the‐art lipid‐based pharmaceutical drug delivery system. Advantages of SLNs include high biocompatibility, low immunogenicity, superiority in drug encapsulation capacity, and improved colloidal stability. They became widely known in late 2020, as several COVID‐19 vaccines are built upon SLNs technology. Despite the increasing impact, the characterization methods of SLNs are currently very limited especially in biological environment, which hinders fundamental understanding of the delivery mechanism and contributes to relatively low success rate in clinical translations. Here, we present close‐to‐label‐free imaging of deuterated SLNs using the emerging stimulated Raman scattering (SRS) microscopy. The introduction of deuterium to lipid structure renders bioorthogonal chemical specificity. Notably, with this approach, we have achieved ultrahigh single‐particle sensitivity both in vitro and in vivo, even with particle counting ability. Our bioorthogonal chemical imaging modality by SRS microscopy can be generalized to visualize a wide spectrum of lipid‐based drug carriers with high spatiotemporal resolution, chemical specificity, and ultimate sensitivity. This work opens up ways to address critical questions in SLN drug delivery and could also facilitate innovations in lipid nanotechnology and clinical translations.

Key points

Direct imaging of lipid nanocarrier that is the basis of the COVID‐19 vaccines.

Novel single‐particle imaging technique applied to nanomedicine.

Frontier of nanomaterial, optical imaging, and molecular pharmacology.

INTRODUCTION

Nanomedicine holds great potential as a novel drug delivery platform.¹ Compared to traditional methods, nanomedicine shows improved delivery efficiency, decreased toxicity, and much higher versatility for a great variety of cargos.^2,3 As such, nanomedicine is expected to revolutionize modern pharmaceutical industry.⁴ Among the key members of nanomedicine, solid lipid nanoparticles (SLNs) are a type of novel lipid-based pharmaceutical formulation and are considered promising in improving drug delivery efficiency.⁵ The popularity of SLNs has been greatly promoted by the advent of COVID-19 vaccines which owe their success to SLNs that took decades to refine. In addition, several RNA-encapsulated SLN formulations are currently undergoing FDA clinical trial.⁶

Generally speaking, SLNs consist of a solid lipid core surrounded by surfactants and cosurfactants to improve their colloidal stability.⁷ Such versatile core–shell structure allows for specialized transportation of both lipophilic and hydrophilic drugs to enhance bioavailability and reduce toxicity.^7,8 SLNs exhibit several advantages that make them promising nanomedicine candidates. First, SLNs are biocompatible and easily biodegradable. As almost all components of SLNs can be made bio-native, low immunogenicity is present at the nano-bio interface.⁵ Second, the lipid core allows efficient delivery of lipophilic small molecule drugs with high bioavailability and low toxicity.⁹ Alternatively, the use of phospholipids and ionizable lipids promotes the delivery of hydrophilic drugs.¹⁰ Third, the formulation of SLNs can be modified to achieve different drug-delivering parameters, including bioavailability, half-life, and diffusing rate. These can be performed through careful tuning of size, surface charge, lipid composition, molecular weight, length of PEGylation, and so on.^11,12 Compared to preexisting carriers such as liposome and niosomes, SLNs exhibit the advantage of enhanced encapsulation efficiency for hydrophobic drugs, long colloidal stability, excellent reproducibility, and low drug leakage.⁶

Despite tremendous progress in the development and commercialization, SLNs share the common frustration faced by the entire nanomedicine field: relatively low success rate in clinical translations.¹³ In fact, among the thousands of clinical trials involving nanomedicine, only a few have passed regulatory clearance and went on market. Unrepresentative model systems, lack of rational drug carrier design, and the knowledge gap of bio-nano interactions are among the most common factors behind this phenomenon.^4,14,15 From a perspective of measurement science, the knowledge gap of bio-nano interactions is largely due to the lack of suitable physical methods to characterize nanocarriers in vivo.¹⁶ Without knowing the precise behaviors of SLNs in vivo, one has to rely on a black-box type of engineering approach which is bound to be less effective.

Characterization methods of SLNs are very limited currently. One of the most commonly used techniques is electron microscopy, including scanning electron microscopy, transmission electron microscopy, and cryo-transmission electron microscopy.¹⁷ However, the lack of chemical specificity limits their detection of SLNs inside the biological matrix. Mass-based techniques show compelling capability in vitro, but the lack of distinct chemical characteristics in SLNs poses tremendous challenges in detecting SLNs in vivo.¹⁸ Most of the characterization of SLNs in biological environments relies on fluorescent microscopy. Generally, lipophilic fluorescent dye or drug is incorporated into SLNs during nanoparticle synthesis, and the nanoparticle is subsequently tracked through the fluorescent signal.¹⁹ This technique, however, possesses intrinsic limitations. As SLNs are designed to distribute cargo inside biological tissues, fluorescent dyes or drugs will eventually leak out, leading to nonspecific false positive signals.^20,21 This creates a huge hurdle for unambiguous detecting SLNs in vivo. Furthermore, fluorescence signal suffers from photobleaching, tissue auto-fluorescence, and self-quenching, making quantification of SLNs nearly impossible.

Here, we report direct imaging of SLNs (∼150 nm in diameter) with minimal labeling. Our technique is built on the emerging stimulated Raman scattering (SRS) microscopy.^22,23 To our best knowledge, this is the first time that Raman microscopy is being employed for imaging SLNs in biological environment. To render bioorthogonal chemical specificity, we have introduced deuterium isotope to the lipid structure. When coupled with highly sensitive SRS microscopy, we have achieved ultrahigh single-particle sensitivity both in vitro and in vivo, even with particle counting ability. Our bioorthogonal chemical imaging strategy of the nanocarrier itself could provide valuable information about the in vivo behaviors of SLNs, thus paving the way for rational nanomedicine design in the future.

METHODS AND EXPERIMENT RESULTS

Stimulated Raman scattering (SRS): principles and instrumentation

Raman imaging is commonly used for the label-free detection of macromolecules due to the inherent chemical specificity, reduced water background, and high spatial resolution. However, the extremely low spontaneous Raman scattering cross section of vibrational mode (∼10⁻³⁰ cm²) has set a fundamental bottleneck for the achievable imaging speed and sensitivity, thus limiting its utility in nanomedicine studies. So far, to our knowledge, there has been no report of direct Raman imaging of a single 100-nm lipid nanoparticle in vivo.¹⁸

SRS microscopy is an emerging vibrational imaging technique that overcomes the drawbacks of conventional Raman by quantum signal amplification through stimulated emission.²² An enhancement factor of 10⁸ in Raman transition rate can be achieved based on previous calculations.²³ This enormous gain translates to much improved detection sensitivity of SRS and more than 10³ folds faster imaging speed, compared to the conventional spontaneous Raman microscopy. Moreover, SRS is intrinsically compatible with other common imaging modalities such as fluorescence microscopy, adding to the utility through multimodal imaging.²⁴ Hence, we aim to employ SRS microscopy to image SLNs in biological environment.

The SRS instrumentation used in this study is the same as described before²⁴ (Figure 1a). In brief, an integrated laser system (Applied Physics and Electronics Inc., picoEMERALD) consisting of a tunable pump beam (720–990 nm, 5−6 ps) and a fixed Stokes beam (1064 nm, 6 ps) modulated by an electro-optic modulator (at 8 MHz) were coupled into a commercial confocal microscope (Olympus, FV1200). The two synchronized beams were spatiotemporally overlapped and then focused onto the sample for laser scanning through a 25X microscope objective (Olympus XLPlan N 1.05 N.A. MP) and collected by a high numerical aperture (N.A. = 1.4) oil immersion condenser. The stimulated Raman loss signal of the pump beam was detected through a large-area photodiode and demodulated by a high-frequency lock-in amplifier (HF2LI, Zurich instrument). Images were acquired by Olympus software and processed using ImageJ and MATLAB. All images were acquired at 100 mW pump power and 150 mW stokes power on sample with a pixel dwell time of 4 µs.

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Synthesis and spectroscopy of deuterated solid lipid nanoparticles (D-SLNs)

Raman spectroscopy of SLNs with C–C, C–H, and fingerprint features has been reported outside biological matrix.^25–27 However, these strict label-free techniques encounter the same challenge faced by electron microscopy and mass-based imaging techniques: lack of bioorthogonality. The internal C–H vibrational modes from cellular lipids and proteins strongly interfere with SLN signal. Besides, some C–H feature also overlaps with Raman peak of H₂O background, and hence, their vibration signal is easily overwhelmed.

To solve the bioorthogonality issue, our idea is to introduce deuterium-labeled fatty acid as a bio-native, yet bioorthogonal compound into the synthesis of SLNs. Carbon–deuterium (C–D) bond displays a distinctive strong broadband peak (∼2100 cm⁻¹) in the cell-silent region (1800–2900 cm⁻¹). Compared to the original C–H peak, C–D Raman signal is free from cellular interference and water background. The introduction of deuterium replacing hydrogen in C–H bonds can be considered minimal labeling, whose effect on biochemical interactions of small molecules can be mostly neglected.²⁸ There have been multiple applications on utilizing C–D bonds for in vivo detection of small molecule metabolism, water metabolism at both the cellular and animal level.^29–31 Thus, it is reasonable to assume that the deuterated-SLNs (D-SLNs) will behave similarly to the regular SLNs in biological contexts, making D-SLNs a perfect substitute to study the behavior of SLNs in vivo.

The synthesis of D-SLNs was adopted and slightly modified from previous protocols³² (Figure 1b,c). Briefly, lecithin, 1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy(polyethylene glycol)-2000] (ammonium salt) (DSPE-PEG-2000, Avanti Polar Lipids, 474922-82-2, MFCD02262246), deuterated stearic acid (d₃₅-SA, Sigma-Aldrich 448249) with a mass ratio of 5:1:10 were dissolved in 0.5 mL dichloromethane and mixed with 5 mL 0.1% poloxamer 188 solution (Thermo Scientific, J6608736). This formulation can be considered a proper model of SLNs, as it contains all key components, including the PEGylated phospholipid. The mixture was then homogenized using a probe sonicator (50% amplitude, 40 s) and evaporated at 65°C for 15 min to remove excess solvent and continued under room temperature for 1 h. The resulting emulsion was filtered using 0.22 µm pore filters and stored at 4°C. A transparent, homogenous solution was obtained through this synthesis method (Figure 1d), indicating the colloidal stability of synthesized D-SLNs.

The size distribution of the D-SLNs was measured by dynamic light scattering (DLS, Malvern Zetasizer, nano-ZS), with the peak at ∼150 nm diameter and a standard deviation of 38 nm. Regular SLNs were synthesized with the same method with non-isotope stearic acid. The spontaneous Raman spectra of SLNs, D-SLNs, and deuterated stearic acid (solid at room temperature) are shown in Figure 1e. Compared to regular SLNs, D-SLNs exhibit expected shifted C–D features in the cell-silent region, with a spectral shape resembling that of deuterated stearic acid, which confirms the deuterated solid core inside D-SLNs. A narrow and prominent peak at 2100 cm⁻¹ features the compact arrangement of C–D bonds in the solid state. The residue C–H peak in the spectrum of D-SLNs may come from other non-deuterated components of the D-SLN such as lecithin and DSPE-PEG. To our best knowledge, this is the first report of D-SLNs.

In vitro measurement: D-SLNs imaging with single-particle sensitivity

We now test SRS imaging of D-SLNs in vitro. To quantify the SRS intensity achievable from a single particle, a diluted aliquot of D-SLNs was mixed with 4% agarose gel to immobilize particles for in vitro study (Figure 2). As shown in Figure 2a,b, the SRS images were acquired at 2100 and 2000 cm⁻¹, corresponding to the on- and off-resonance condition, respectively. Particle-like signals in the C–D channel can be readily observed across the field of view, and the signal can be completely tuned off by a ∼100 cm⁻¹ shift. A high signal-to-noise ratio (SNR) of 50 or higher was detected for single particle, indicating an ultrahigh detection sensitivity.

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The maximum intensity value of each particle was then recorded. Upon statistical analysis of the particle signals, semi-discrete distribution of signals can be observed from the histogram (Figure 2c). The quantized intensity level in the histogram suggests the existence of the “minimal” single particles and their aggregations, which confirmed the single-particle sensitivity of our technique. To further validate single-particle signal, we conducted a numerical estimation of the signal based on C–H stretching and C–O Raman intensities reported previously.^24,33 Assuming the smallest signal we observed comes from a 90-nm-diameter spherical solid core consisting of pure stearic acid (M.W. 284.48 g/mol) with a density of 941 kg/m³, we can calculate that there are approximately 12 million CH₂ units in such a particle. The signal obtained from a single CH₂ was estimated to be 2.5 times C–O bond, thus 1/20 of the EdU alkyne signal. We can then estimate the detectable SRL signal from a single particle to be ΔI/I = 1 × 10⁻⁵. This should give us an SNR of ∼50 as the noise from instrument was measured to be 2 × 10⁻⁷. This estimation is in good agreement with our experiment results, further supporting the ability to detect single-particle signals.

The particle counting capability is demonstrated in a zoom-in view (Figure 2d,e), where quantized signals of individual clusters are plotted, representing the intensity level ranging from single particle to tetramer. The full-width-at-half-maximum of the smallest particle was measured to be around ∼450 nm, which is consistent with the instrument resolution limit. To confirm, the signal is indeed from synthesized deuterated lipid nanoparticles, we also acquired a spectral image of D-SLNs (Figure 2f,g). Each individual particle can be gradually tuned on and then tuned off when sweeping across different pump wavelengths. The exhibited spectral shape corresponds perfectly with the bulk material measurements, ruling out the possibility of artifacts and contaminations during the experiment. Unlike fluorescent imaging, no significant photobleaching of signals was observed under the SRS imaging condition across 60 frames, which allows for robust quantitative imaging and particle tracking capabilities (Figure 2h).

SRS imaging of single D-SLN uptake in cellular environments

We next demonstrate bioorthogonal imaging of D-SLNs inside complex biological environments. Macrophage cells are one of the most common model systems used in nanomedicine research because they mimic the native immune response of organisms.^34,35 Detailed study of the interaction of macrophage cells and SLNs could provide information about the immunogenicity of lipid nanoparticles, as well as the entry and degradation pathways inside cells. In short, RAW264.7 macrophage cells were incubated with the growth media containing D-SLNs aliquot synthesized in the abovementioned method. The cells were then washed with PBS three times to remove any excess D-SLNs, fixed with 4% PFA, and mounted for SRS imaging.

A significant amount of nanoparticles were taken up by macrophages after 24 h, as shown in Figure 3. Most particles were observed to accumulate in the cell cytoplasm rather than residing inside the cell nucleus. This supports the in vivo efficiency of the formulation we adopted for SLNs. Interestingly, cells that were undergoing mitosis seemed to take up more nanoparticles than others. Thanks to the bioorthogonality of the introduced deuterium isotope, D-SLNs can be successfully detected in the C–D on-channel at 2100 cm⁻¹ without interference from endogenous biomolecules. The signal can be nicely tuned off in the off-resonant channel at 2000 cm⁻¹. The authenticity of particle identity was further confirmed through spectral imaging, as shown in Figure 3e. The spectral profile is analogs to the reported spectra in the previous session in vitro, with a sharp signature peak at 2100 cm⁻¹. Single D-SLN was identified inside macrophage cells (Figure 3f,g), according to the predefined value of single-particle intensity. Four diffraction-limited spots with intensities corresponding to single particle were highlighted, validating our capability of detecting individual particles in vivo.

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A more careful interrogation of the images revealed several findings. First, small clusters of D-SLNs were observed in several cells, which raises the question of whether D-SLNs aggregate inside cell organelles due to changes of local pH conditions. Second, the C–D on-resonance image shares resembling features with the C–H channel. This might be explained by the residue C–H bonds from cosurfactants in lipid nanoparticles. Another possibility is that D-SLN might have induced local lipid droplet formation or simply fused with internal droplets. Third, the detected C–D peak is slightly broader than the previous result from gel measurements (Figure 2), which could stem from the local physiochemical environment changes and increase in cross-phase background. An alternative explanation would be the partial degradation and protein coronation by the cell has altered the composition of the D-SLN core and was causing spectral variations in the C–D profile. Finally, the cell body also presents some faint C–D positive signal as well. This could be explained by the partial degradation of some D-SLNs and cell's utilization of d₃₅-stearic acid to produce isotope-labeled biological matter. These open questions deserve future investigations.

DISCUSSION AND CONCLUSION

In summary, we have synthesized deuterated SLNs and demonstrated the SRS spectral imaging capability down to single-particle level with high spatial resolution in biological environments. We emphasize several technical points here on achieving such a superb detection sensitivity. First, we chose C–D as Raman-active probe to label our SLN model system with minimal perturbation. The enrichment of C–H/C–D chemical bonds inside lipid-based nanoparticle core raises the local concentration of Raman-active bonds, hence amplifying the Raman signal. Second, the solid-phase core formed by long-chain fatty acid with high melting temperature exhibits higher conformational order of the acyl chain, making the C–H/C–D vibrational feature more spectrally intense (i.e., narrower Raman peak), compared to other formulation such as lipid nano-emulsion. Third, SRS imaging technique offers orders of magnitude signal enhancement from stimulated emission compared to spontaneous Raman process. Furthermore, SRS signal is linearly proportional to molecular concentration, enabling the qualitative particle counting in our study. To our knowledge, this is the first time that single-particle Raman imaging is achieved on lipid-based nanomedicine inside biological context.

It is worth noting to point out that the SLNs synthesized in this report are merely a model system. Our use of deuterated lipids to label lipid-based nanoparticles for bioorthogonal SRS imaging in vivo is a generalizable strategy. An SNR of over 100:1 was observed with 100% substituted stearic acid solid core, which indicates that a composition of 1% deuterated lipids in a lipid nanoparticle could give an SNR∼1 and will be theoretically observable under current instrumentation. Thus, this strategy can be applied to all types of lipid-containing nanomedicine members, including liposome, noisome, and the state-of-the-art lipid-structure nanoparticles,³⁶ and is compatible with systems of different surfaces, charges, sizes, and lipid constructions.

Our technique allows for the direct observation of lipid nanocarriers themselves (rather than the cargo) and reveals the true biodistribution of them in cells and tissues with ultimate resolution and sensitivity. This could open the door for mechanistic studies of important problems in the field of nanomedicine. For example, the myth of how SLNs are distributed in different tissues and cell types and degraded after they enter the human body is still elusive, despite the ubiquitous use of SLN-based COVID-19 vaccines since the late 2020. We believe that the technique provided in this report could illuminate the science and technology of lipid-based nanomedicine, in particular, by providing insight as to how to improve targeted delivery efficiency and facilitate rational nanocarrier design.

AUTHOR CONTRIBUTIONS

Xiaoqi Lang and Wei Min designed the experiments with the input from Xin Gao, Mian Wei, and Naixin Qian. Xiaoqi Lang prepared the samples, conducted the experiments, and analyzed the data with the help of Xin Gao, Mian Wei, and Naixin Qian. Xin Gao, Xiaoqi Lang, and Wei Min wrote the main paper with the input from all coauthors.

ACKNOWLEDGMENTS

We thank Dr. Donghui Song and Dr. Chenyi Mao for the assistance on sample preparation and discussion. We thank support from NIH (R01 GM132860 and R01 EB029523).

CONFLICT OF INTEREST STATEMENT

The authors declare no conflict of interests.

DATA AVAILABILITY STATEMENT

All raw images and data are available from the corresponding author upon request.

ETHICS STATEMENT

The authors confirm the material is the author's own original work, and has not been published elsewhere. The authors are committed to upholding ethical standards in research and ensuring the accuracy and reliability of the research data through rigorous experimental design, data analysis, and reporting. All data reported in this study were collected in a transparent manner, and any potential conflicts of interest were disclosed and addressed appropriately.

PEER REVIEW

The peer review history for this article is available at .

References

Huang X, Kong N, Zhang X, Cao Y, Langer R, Tao W. The landscape of mRNA nanomedicine. Nat Med. 2022;28:2273‐2287.

Quader S, Kataoka K, Cabral H. Nanomedicine for brain cancer. Adv Drug Deliv Rev. 2022;182: [eLocator: 114115].

Bhatia SN, Chen X, Dobrovolskaia MA, Lammers T. Cancer nanomedicine. Nat Rev Cancer. 2022;22:550‐556.

Rasool M, Malik A, Waquar S, et al. New challenges in the use of nanomedicine in cancer therapy. Bioengineered. 2022;13:759‐773.

Duan Y, Dhar A, Patel C, et al. A brief review on solid lipid nanoparticles: part and parcel of contemporary drug delivery systems. RSC Adv. 2020;10:26777‐26791.

Thi TTH, Suys EJ, Lee JS, Nguyen DH, Park KD, Truong NP. Lipid‐based nanoparticles in the clinic and clinical trials: from cancer nanomedicine to COVID‐19 vaccines. Vaccines. 2021;9:359.

Moraes‐Lacerda T, de Jesus MB. Mechanisms of solid lipid nanoparticles‐triggered signaling pathways in eukaryotic cells. Colloids Surf B Biointerfaces. 2022;220: [eLocator: 112863].

Mendes BB, Conniot J, Avital A, et al. Nanodelivery of nucleic acids. Nat Rev Methods Primers. 2022;2:1‐21.

Harde H, Das M, Jain S. Solid lipid nanoparticles: an oral bioavailability enhancer vehicle. Expert Opin Drug Deliv. 2011;8:1407‐1424.

Mirchandani Y, Patravale VB, S B. Solid lipid nanoparticles for hydrophilic drugs. J Control Release. 2021;335:457‐464.

Xu Y, Fourniols T, Labrak Y, Préat V, Beloqui A, des Rieux A. Surface modification of lipid‐based nanoparticles. ACS Nano. 2022;16:7168‐7196.

Vitorino C, Carvalho FA, Almeida AJ, Sousa JJ, Pais AACC. The size of solid lipid nanoparticles: an interpretation from experimental design. Colloids Surf B Biointerfaces. 2011;84:117‐130.

van der Meel R, Sulheim E, Shi Y, Kiessling F, Mulder WJ, Lammers T. Smart cancer nanomedicine. Nat Nanotechnol. 2019;14:1007‐1017.

Sun D, Zhou S, Gao W. What went wrong with anticancer nanomedicine design and how to make it right. ACS Nano. 2020;14:12281‐12290.

Björnmalm M, Thurecht KJ, Michael M, Scott AM, Caruso F. Bridging bio‐nano science and cancer nanomedicine. ACS Nano. 2017;11:9594‐9613.

Li P, Wang D, Hu J, Yang X. The role of imaging in targeted delivery of nanomedicine for cancer therapy. Adv Drug Deliv Rev. 2022;189: [eLocator: 114447].

Gordillo‐Galeano A, Mora‐Huertas CE. Solid lipid nanoparticles and nanostructured lipid carriers: a review emphasizing on particle structure and drug release. Eur J Pharm Biopharm. 2018;133:285‐308.

Fan Y, Marioli M, Zhang K. Analytical characterization of liposomes and other lipid nanoparticles for drug delivery. J Pharm Biomed Anal. 2021;192: [eLocator: 113642].

Barbosa HFG, Piva HL, Tedesco AC. Lysosomal tracking with red‐fluorescent solid lipid nanoparticles. OpenNano. 2022;8: [eLocator: 100069].

Akhoond Zardini A, Mohebbi M, Farhoosh R, Bolurian S. Production and characterization of nanostructured lipid carriers and solid lipid nanoparticles containing lycopene for food fortification. J Food Sci Technol. 2018;55:287‐298.

Shu L, Fu F, Huang Z, Huang Y, Hu P, Pan X. Nanostructure of DiR‐loaded solid lipid nanoparticles with potential bioimaging functions. AAPS PharmSciTech. 2020;21:321.

Freudiger CW, Min W, Saar BG, et al. Label‐free biomedical imaging with high sensitivity by stimulated Raman scattering microscopy. Science. 2008;322:1857‐1861.

Min W, Freudiger CW, Lu S, Xie XS. Coherent nonlinear optical imaging: beyond fluorescence microscopy. Annu Rev Phys Chem. 2011;62:507‐530.

Wei L, Chen Z, Shi L, et al. Super‐multiplex vibrational imaging. Nature. 2017;544:465‐470.

Jores K, Haberland A, Wartewig S, Mäder K, Mehnert W. Solid lipid nanoparticles (SLN) and oil‐loaded SLN studied by spectrofluorometry and Raman spectroscopy. Pharm Res. 2005;22:1887‐1897.

Cherney DP, Conboy JC, Harris JM. Optical‐trapping Raman microscopy detection of single unilamellar lipid vesicles. Anal Chem. 2003;75:6621‐6628.

De Gaetano F, Cristiano MC, Venuti V, et al. Rutin‐loaded solid lipid nanoparticles: characterization and in vitro evaluation. Molecules. 2021;26:1039.

Shen Y, Hu F, Min W. Raman imaging of small biomolecules. Annu Rev Biophys. 2019;48:347‐369.

Hu F, Lamprecht MR, Wei L, Morrison B, Min W. Bioorthogonal chemical imaging of metabolic activities in live mammalian hippocampal tissues with stimulated Raman scattering. Sci Rep. 2016;6:1‐11.

Shi L, Zheng C, Shen Y, et al. Optical imaging of metabolic dynamics in animals. Nat Commun. 2018;9:2995.

Zhang L, Shi L, Shen Y, et al. Spectral tracing of deuterium for imaging glucose metabolism. Nat Biomed Eng. 2019;3:402‐413.

Naguib YW, Rodriguez BL, Li X, et al. Solid lipid nanoparticle formulations of docetaxel prepared with high melting point triglycerides: in vitro and in vivo evaluation. Mol Pharm. 2014;11:1239‐1249.

Zhao Z, Chen C, Wei S, et al. Ultra‐bright Raman dots for multiplexed optical imaging. Nat Commun. 2021;12:1305.

Chen W, Schilperoort M, Cao Y, Shi J, Tabas I, Tao W. Macrophage‐targeted nanomedicine for the diagnosis and treatment of atherosclerosis. Nat Rev Cardiol. 2022;19:228‐249.

Ma C, Wu M, Ye W, et al. Inhalable solid lipid nanoparticles for intracellular tuberculosis infection therapy: macrophage‐targeting and pH‐sensitive properties. Drug Deliv Transl Res. 2021;11:1218‐1235.

Mahmoud K, Swidan S, El‐Nabarawi M, Teaima M. Lipid based nanoparticles as a novel treatment modality for hepatocellular carcinoma: a comprehensive review on targeting and recent advances. J Nanobiotechnol. 2022;20:109.