High-resolution sub-cellular imaging has greatly advanced the cell biology field, making it possible to visualize organelles and other critical sub-cellular structures and molecular assemblies (DʼEste et al., 2024; Lučić et al., 2013). For example, the integration of high-resolution techniques like multiphoton microscopy (MPM) has shown promising diagnostic accuracy in assessing pediatric tissues and tumors (Goedeke et al., 2019). This has enhanced our understanding of normal developmental processes, but has also allowed assessment of cellular changes during disease development and progression (Mojahed et al., 2022; Hayes and Melrose, 2023).

The field of tissue engineering has made significant advancements in replicating the complex structure and function of biological tissues. Using these engineered tissues to mimic 3D complex human tissues has allowed researchers to obtain a more accurate understanding of disease mechanisms, whilst also enabling screening of pharmaceuticals to assess drug efficacy and toxicity. In the cardiovascular field, 3D engineered heart tissues have allowed researchers to study cell-cell and cell-ECM (extracellular matrix) interactions that cannot be studied in 2D monocultures (Simmons et al., 2024; Van Spreeuwel et al., 2014; Thavandiran et al., 2013; Michas et al., 2022; Ma et al., 2023). For example, in a recent study from our team, we found that induced pluripotent stem cell (iPSC) derived cardiomyocytes with a hypertrophic cardiomyopathy (HCM) associated genotype exhibited structural, pharmacologic and physiologic hallmarks of HCM in 3D engineered tissues that we were unable to appreciate in a traditional 2D culture (Guo et al., 2024). Analogously, in studying arrhythmogenic cardiomyopathy (ACM), a disease that has been linked to contractile deficits in vivo, (Chen et al., 2024) observed that isolated iPSC-cardiomyocytes with an ACM-linked genotype were largely similar to (if not more contractile than) isogenic controls. However, growing the iPSC-cardiomyocytes with an ACM genotype in 3D engineered tissues unveiled deficits in contractility that were attributed to weaker cell-cell mechanical coupling (Zhang et al., 2021).

In cancer research, 3D tumor models provide a realistic environment for studying tumor progression and testing anticancer drugs (Kang et al., 2020). Drug discovery and toxicology also benefit from 3D engineered tissues, since they offer more accurate models for evaluating drug efficiency and safety compared to traditional 2D cell cultures (Khetani and Bhatia, 2008; Khalil et al., 2020; Simmons et al., 2022). For example, a 3D tumor spheroid model incorporating stromal fibroblasts has been developed to better mimic the in vivo tumor microenvironment. This model not only enables the study of tumor-stroma interactions and the role of cancer-associated fibroblasts (CAF) in cancer progression but also facilitates drug discovery (Shao et al., 2020).

Besides their translational role in disease modeling and drug development, engineered tissues have also allowed scientists to study fundamental questions in developmental biology (including tissue morphogenesis and cell differentiation; (Zuncheddu et al., 2021; Pien et al., 2023)), and biophysics [including the mechanical properties and behaviors of various tissues under different physiological conditions; (Guo et al., 2021a; Li et al., 2023)]. The development of organoid technology, in which “organ-like” structures develop from progenitor and/or stem cells in vitro to recapitulate key aspects of normal tissue development, has allowed scientists to study “organs in a dish” that replicate the phenotypic and functional properties of real tissue environments (Moysidou et al., 2021; Bose et al., 2019; Bhatia and Ingber, 2014). Importantly, organoids allow in vitro differentiation of cell types (notably, certain endodermal cell populations) that have historically proven challenging to derive through directed differentiation methods (Vunjak-Novakovic et al., 2021; Hofer and Lutolf, 2021).

Given that cells exhibit more “in vivo-like” signaling, pharmacology and behavior within 3D engineered tissues, organoids and other complex in vitro assembled structures compared to what they would exhibit in more “standard” 2D monoculture environments, a longstanding question for bioengineers and cell biologists is “how are cells sensing their environment?.” Forces that cells experience are dramatically different in 3D tissues as opposed to standard 2D culture, and indeed, recent studies by our team and others, suggest that mechanical stretch affects electrophysiology of engineered heart muscle (Simmons et al., 2024; Guo and Huebsch, 2020; Wang C. et al., 2023; Schuftan et al., 2024; Boudou et al., 2012; DePalma et al., 2023; Abilez et al., 2018; Tulloch et al., 2011; Huebsch, 2019; Guo et al., 2021b). This may be linked to either improved expression and/or intracellular transport of ion channels (Simmons et al., 2024; Marchal et al., 2021). In a broader context, Chaudhuri et al. provided a comprehensive review of the complex mechanical properties of tissue and ECMs including viscoelasticity, viscoplasticity, and nonlinear elasticity, and discussed how these properties affect cellular behaviors (Chaudhuri et al., 2020). Adding to this, Gong et al. found that maximum cell spreading occurs at an optimal viscosity level on soft substrates for low ECM rigidity, where the substrate relaxation time falls between clutch binding and lifetime timescales, enhancing cell-ECM adhesion, while on stiff substrates, viscosity has no effect due to clutch saturation, which provides insights for designing biomaterials to optimize cell adhesion and mechano-sensing (Gong et al., 2018). Moreover, in 3D cultures, cancer cells exhibit enhanced drug resistance mechanisms compared to 2D cultures, this is attributed to the more realistic cell-cell and cell-matrix interactions in 3D environments, which affect gene expression profiles and signaling pathways related to drug metabolism and resistance. Hypoxic conditions and nutrient gradients present in 3D tumor spheroids activate hypoxia-inducible factors (HIFs), leading to the upregulation of drug resistance genes and proteins (Bloise et al., 2024; Atat et al., 2022). Additionally, cancer cells in 3D environments use invadopodia to sense and dynamically respond to the mechanical properties of the ECM, such as its stiffness and plasticity. Through chemo-mechanical feedback, invadopodia can adjust their growth patterns, enabling cancer cells to invade tissues more effectively through chemo-mechanical signaling feedback loops (Gong et al., 2021). Therefore, understanding the molecular mechanisms cells use to sense their 3D environment will allow bioengineers to develop better in vitro model systems and may provide molecular targets that can be directly exploited for therapy.

A challenge in unveiling these sensing mechanisms is that such studies will require detailed analysis of subcellular architecture. This presents a fundamental technical challenge: these environmental changes that provoke a more “in vivo-like phenotype” requires putting cells into 3D, whereas traditionally, detailed analysis of subcellular architecture has required growing cells in 2D environments (e.g., culture on glass coverslips). While artificial, this culture approach places cells very close (typically, within 150 μm) to microscope objectives, allowing researchers to use high-resolution optics with high numerical-aperture immersion media (e.g., oil) and in turn facilitating robust analysis of sub-cellular architecture (Zuncheddu et al., 2021; Pien et al., 2023; Berry et al., 2021; Zhao et al., 2022; O’Connor et al., 2022; Tan et al., 2004).

Advancements in optics, coupled with advancements in the fluorescent probes used for imaging, offer promising means to combine high resolution subcellular imaging with physiologically relevant engineered tissues and organoids (Li et al., 2019; Dekkers et al., 2019; Hilzenrat et al., 2022; Wang S. et al., 2023). To achieve a high-resolution image, it is essential to balance spatial and temporal resolution, maintain an optimal signal-to-noise ratio, and avoid photobleaching (Figure 1). Here, we first review basic physical principles underlying fluorescence imaging. We then discuss promising approaches that tissue engineers have taken to enable high-resolution, sub-cellular imaging of 3D tissues, and provide examples of some of the key insights gained from these analyses. The lateral, axial, and temporal resolutions of these methods, along with their imaging depths, are detailed in Table 1, which also includes schematics of the focal plane for each technique. We aim to simplify the decision-making process for researchers choosing the most optimal imaging approach by presenting simplified guidance on choosing methods based on specific research needs. Excellent reviews of imaging methods, which delve more deeply into the physics underlying the various methods we discuss, are available elsewhere (Huang et al., 2009; Winter and Shroff, 2014; Leung and Chou, 2011). Another key point of our review is that while most high-resolution imaging methods focus on 2D samples or sectioned 3D tissue, we specifically highlight methods of whole-mount staining and optical sectioning for 3D tissues, which allows for better imaging of intact 3D structures, an area that remains underrepresented in the current literature.

Fluorescence microscopy is a cornerstone technique in biological sciences, enabling the visualization of cellular structures and functions by exploiting the principles of fluorescence (Masters, 2014). The underlying mechanism involves the excitation of fluorophores, molecules that can re-emit light upon excitation that is specifically attached to target molecules within the sample, such as proteins or nucleic acids. When exposed to light of a specific wavelength (typically from a UV source like a mercury arc lamp), these fluorophores absorb photons, exciting electrons to a higher energy state. The electrons, inherently unstable at this level, quickly return to their ground state, releasing photons with a longer wavelength and lower energy than the absorbed light (Sanderson et al., 2014). The change in wavelength between the photons that excite fluorescence and the photons emitted is called the Stokes shift (Peng et al., 2005). The Stokes shift occurs because the emitted light is of lower energy due to losses during the nonradiative transitions between excited and ground states, a process that Jablonski energy diagrams explains (Figure 2A) (Yao et al., 2014; Berezin and Achilefu, 2010). Detailed understanding of the physics of decay of the higher-energy states has allowed researchers to control Stokes shift and exploit non-radiative methods of energy decay (e.g., Förster Resonance Energy Transfer, FRET) for analyzing sub-cellular architecture (detailed in Section 3).

In fluorescence microscopy, optical filters and epi-illumination techniques are used to enhance quality of the images. An excitation filter selects the optimal wavelength to excite the fluorophore. A barrier filter blocks excitation wavelengths while passing longer emission wavelengths, and a dichroic mirror directs excitation light to the sample and emitted fluorescence to the detector. This allows the objective lens to both illuminate the specimen with the excitation light and to collect the emitted fluorescence (Figure 3A) (Lichtman and Conchello, 2005; Combs and Shroff, 2017). In diffraction limited (e.g., not super-resolution) optical systems, the minimum distance d between objects that can be resolved is related to the wavelength of light (λ) and the numerical aperture (N.A.) is dictated by the equation d = λ 2 N . A .

Within the visible wavelength range, this sets a diffraction limit near 250–300 nm (Thorne and Blandford, 2017). This is significantly smaller than a mammalian cell (10 s of μm) but large compared to proteins (10 s of nm). Lower wavelength, higher energy photons (e.g., x-rays) yield a smaller diffraction limit that can allow resolving protein-protein interactions, but this often limits the ability to image live samples.

Atop the diffraction limit, epifluorescence microscopy methods face additional limitations in resolution and contrast due to the full-sample illumination strategy (Bates et al., 2008). While in theory the resolution limit of epifluorescence imaging is about 250 nm laterally and 500 nm axially (Lidke and Lidke, 2012), in practice it is often challenging to separate objects closer than 1 μm. The Point Spread Function (PSF) describes how light spreads as it passes through the microscope optics (Cole et al., 2011; Pankajakshan et al., 2009). In epifluorescence, there is high background intensity from out-of-focus fluorescence, limiting the clarity of the images (Figure 2B) (Huang et al., 2009; Sung et al., 2002; Huang et al., 2008a). These challenges have spurred the development of advanced microscopy techniques that enhance image quality by manipulating the light path and minimizing unwanted fluorescence (Heilemann, 2010; Hell, 2003; Hell, 2009; Helmchen and Denk, 2002). Such improvements not only reduce photobleaching effects, but also increase the throughput and specificity of fluorescence imaging systems, which allows for more detailed and quantitative studies of live cells and tissue (White, 2005; Ishikawa-Ankerhold et al., 2012). As researchers continue to refine these methods, the ability to visualize biological processes with greater precision enhances our understanding of complex cellular mechanisms within biological samples such as 3D engineered tissues.

In the prior section, we discussed imaging innovations which center on controlling sample illumination and emitted photon detection. A second, parallel means for enhancing spatiotemporal resolution exploits an ever-growing body of knowledge of photo-physics, which details how materials respond to light. Changes in the local environment of an electron can increase the likelihood of the photon-absorption-triggered excited state to decay by non-radiative means (e.g., through pathways besides emission of a lower-energy photon). Because material characteristics (e.g., chemical composition) can be selected to control electrons’ environment in a predictable manner, it has been possible to engineer key fluorescence properties of fluorophores (e.g., quantum yield, Stokes shift) (Lakowicz and Masters, 2008). Here, we detail several imaging modalities that exploit our growing knowledge of photo-physics to explore subcellular structures. A persistent theme in this body of work is that photophysical properties of materials can be exploited to overcome the diffraction limit in resolving nanoscale structures.

In the prior section, we discussed about enhancing spatiotemporal resolution through the exploitation of photophysical properties of materials, which details how materials respond to light. A third method to enhance imaging resolution involves optimizing sample processing. The way a sample is processed can directly affect the image resolution by understanding how light interacts with the tissue. Refining aspects like sample size, transparency, and fixation methods, we enhance light penetration and minimize light scattering. This integration of advance material processing with optical techniques opens a new area for capturing detailed cellular structures with high clarity. Here, we will elaborate on various methods that have been employed to optimize sample processing for better imaging compatibility.

Building upon the advancements in imaging technology, the combination of sophisticated optical techniques, fluorescent probes, and sample preparation techniques has significantly enhanced our ability to explore and visualize subcellular structures in tissue engineering. For instance, Spinning Disc Confocal Microscopy have used a commercially fabricated chromium photomask to enable 3D biological imaging across a wide wavelength range (400–800 nm). This innovation incorporates hundreds to thousands of simultaneous illumination points, reducing peak illumination power density (Halpern et al., 2022). Furthermore, integration of fluorescence lifetime imaging microscopy by an Spinning Disc Confocal Microscopy into a frequency-domain FLIM system, reduces out-of-focus blur and photobleaching while maintaining short acquisition times (Van Munster et al., 2007). In order to get high temporal resolution to study calcium signaling in vascular endothelial cells, Spinning Disc Confocal Microscopy equipped with an electron-multiplying CCD camera has been used to reveal rapid and localized calcium transients which is critical in regulating blood flow and pressure (Nelson et al., 2012). Furthermore, optical photon reassignment (OPR), an advanced super-resolution method using Spinning Disc Confocal Microscopy, have been used to significantly enhance lateral resolution by a factor of 1.37 through a single exposure (Azuma and Kei, 2015). Moreover, to achieve high resolution and contrast in live 3D engineered tissues, a custom-built Spinning Disc Confocal Microscopy with the second near-infrared (NIR-II) optical window have been used to achieve lateral (0.5 ± 0.1 µm) and axial (0.6 ± 0.1 µm) resolution (Zubkovs et al., 2018). Lastly, the combination of engineered tissues with spinning disc confocal microscopy has given us new insights into mitochondrial dynamics. Using Spinning Disc Confocal Microscopy, Ahmadian et al. optimized and quantified live images of 3D mitochondrial networks in mesoangioblasts and mesoangioblast-derived myotubes. This technique reduced background signals and variation in fluorescence intensity, enabling more accurate and reproducible imaging compared to Laser Scanning Confocal Microscopy. This allowed for the identification of key changes in mitochondrial structure and function during cell differentiation, which enhances our understanding of mitochondrial behavior in complex tissue environments (Ahmadian et al., 2024).

Multiple light-sheet microscopy enhances the capabilities of single plane illumination microscopy by reducing photobleaching and enhancing imaging speed and is particularly suitable for imaging. This approach uses spatial filters in the excitation arm to create multiple light-sheets to allow simultaneous illumination of multiple planes within a sample, to facilitate rapid imaging. The multiple light-sheet microscopy system demonstrated the ability to produce thinner light-sheets with increased resolution and reduced crosstalk between the illuminated planes (Mohan et al., 2014). Light-sheet microscopy techniques enhanced by reversible saturable optical fluorescence transitions (RESOLFT) significantly surpass the diffraction limit in axial resolution, enabling 3D imaging of live biological specimens with minimized photodamage and light exposure and it has the ability to produce optical sections up to 5–12 times thinner than those achievable with conventional diffraction limited light-sheet fluorescence microscopy (Hoyer et al., 2016). Advancements in light-sheet fluorescence microscopy have significantly improved high-speed 3D imaging of live tissues, especially for calcium dynamics in cardiac engineered tissues. Light-sheet fluorescence microscopy offers enhanced imaging speed and resolution with minimal light exposure, providing deeper insights into fast physiological processes. Sparks et al. used LSFM to obtain high-resolution images of t-tubule networks in real tissues, revealing their precise 3D structure and organization, which is critical for efficient excitation-contraction coupling. This level of detail is challenging to achieve with laser scanning confocal or two-photon microscopy due to their limitations in depth penetration and resolution (Sparks et al., 2020).

Two-photon microscopy has been used to study the enhancement of the structural and functional properties of engineered heart tissues through chronic electrical stimulation, which is crucial to examine the maturation and functionality of tissue-engineered cardiac constructs. In the study, two-photon microscopy revealed detailed visualization of increased connexin-43 abundance, indicating better gap junction formation and electrical coupling in paced engineered heart tissues, and enabled the measurement of a significantly thicker compact cardiomyocyte layer in paced engineered heart tissue and demonstrated a more homogeneous distribution of cardiomyocytes throughout the tissue depth (Hirt et al., 2014). Two-photon microscopies with near-infrared femtosecond laser pulses have been used to visualize the intratissue elastic fibers within both native and engineered heart valves without the need for invasive procedures such as tissue removal, embedding, or staining. Using multiphoton-induced autofluorescence and second harmonic generation, König et al. could clearly differentiate between elastic fibers and collagenous structures within the extracellular matrix (König et al., 2005). Blazeski et al. used two photon microscopies to visualize and analyze cellular and subcellular structures in engineered heart slices (EHS). This allowed for the precise localization of live cells relative to the matrix and the observation of structural cues promoting cellular alignment, which are crucial for understanding the physiological and pathophysiological roles of the ECM in cardiac tissue engineering (Blazeski et al., 2015). Moreover, to demonstrate that intermittent mechanical straining enhances and accelerates collagen fiber alignment in engineered heart tissues, two. photon microscopy have been used to achieve high resolution in thick 3D engineered tissues (Rubbens et al., 2009). Ye et al. used two-photon laser scanning microscopy to show that SDS best preserves the structural integrity of the 3D engineered heart tissues as decellularization reagent and visualize details about the microstructure of decellularized porcine heart tissue. With this technique, they were able to observe the ultrastructural morphology of type I collagen within the extracellular matrix (ECM) without the need for complicated fixation and washing processes, which can alter the tissue’s natural structure (Ye et al., 2016).

Expanding beyond cardiovascular applications, Chang et al. used third harmonic generation (THG) and two-photon excited fluorescence (2PEF) imaging techniques to non-invasively monitor the functional properties of 3D engineered human adipose tissues to provide detailed, label-free images of lipid droplet formation and cell metabolism over time. THG was used to visualize lipid droplets, while 2PEF assessed the metabolic state of cells through the redox ratio of endogenous FAD and NADH fluorescence. The combination of these imaging approach allowed this team to dynamically correlate lipid accumulation with changes in the metabolic state during adipogenic differentiation. They discovered that cells in adipogenic media showed a significantly lower redox ratio and increased lipid content compared to those in propagation media, providing new insights into the relationship between metabolic changes and lipid biosynthesis in differentiating stem cells (Chang et al., 2013). Meanwhile, Straub et al. used Multifocal Multiphoton Microscopy to achieve non-invasive, high-resolution live cell imaging, which significantly enhanced the understanding of cell dynamics and morphology in three dimensions. This method used an array of high numerical aperture foci for parallel multiphoton excitation, allowing rapid and detailed 3D reconstructions of live cells at speeds up to video rate. This approach is particularly advantageous for dynamic cellular processes that require fast imaging techniques. For instance, they were able to visualize the detailed morphology of neurites and the dynamics of membrane-associated processes, which were not as comprehensively understood with previous imaging technologies due to slower acquisition times and greater photodamage (Straub et al., 2000). To image 3D engineered neuronal networks, a hybrid multiphoton microscope combining scanning-line temporal-focusing with laser-scanning two-photon microscopy have been introduced. This dual approach allows for both rapid volumetric imaging and detailed structural analysis within a single system, which is crucial for investigating dynamic neuronal activities and interactions in 3D cultures. The primary achievement of this method is highly detailed three-dimensional imaging at a microscopic scale across a volume of tissue and at high speeds (tens of volumes per second). This has enabled the observation of over 1,000 developing cells and their complex spontaneous activity patterns within millimeter-scale structures. This system use a regeneratively amplified ultrafast laser to enhance two-photon absorption significantly, which improves the signal-to-noise ratio (Dana et al., 2014).

Super-resolution Stimulated Emission Depletion (STED) microscopy has been used for 2D and 3D cell culture methods to study the molecular organization of apical and lateral membrane domains of epithelial cells. By using an inverted filter mounting strategy, Maraspini et al. were able to gain better access to the apical membrane domains, allowing them to effectively image and resolve the densely packed micro-villi of human enterocytes. Additionally, the optimization of 3D organotypic cell culture enabled the detailed visualization of adhesion complexes in the lateral membrane domain of kidney-derived cells which significantly contributed to our understanding of epithelial membrane organization (Maraspini et al., 2020). Moreover, STED microscopy was utilized to visualize dynamic morphological changes in dendritic spines with a resolution of about 70 nm, significantly enhancing the understanding of synaptic function and plasticity beyond what is possible with confocal microscopy (Nägerl et al., 2008), and STED nanoscopy was used for detailed visualization of mitochondrial dynamics and interactions at high resolution (Liu et al., 2022).

Multicolor three-dimensional stochastic optical reconstruction microscopy (3D STORM) has been used to examine cellular structures and their interactions at nanoscale resolution. By this method Huang et al. were able to image entire mitochondrial networks in mammalian cells, which shows detailed mitochondrial morphologies and their spatial relationships with microtubules that were not visible through epifluorescence microscopy (Huang et al., 2008b). Moreover, STORM was used to examine the nanoscale distribution and clustering characteristics of Angiotensin II Receptor Type 1 (AT1R) on the membrane of PC12 cells. Aldossary et al. found that exposure to hypoxia for 24 h increased the maximum cluster area, suggesting a formation of superclusters. This change in the clustering behavior of AT1Rs under hypoxia might influence the receptor’s function and cell signaling pathways, indicating a potential mechanism by which cells respond to low oxygen levels (Aldossary et al., 2023). Lastly, STORM was used to visualize the endothelial surface glycocalyx (ESG) of cultured endothelial cells, revealing its molecular components and ultrastructure with high spatial resolution (Xia and Fu, 2024).

In this review, we have highlighted the growing importance of high-resolution sub-cellular imaging in the field of tissue engineering. These complex structures offer a more realistic environment for studying cellular behavior and disease compared to traditional 2D cultures. However, obtaining high-resolution images of these dense tissues presents a challenge due to limitations in light penetration and accessibility. Fortunately, advancements in microscopy techniques (like light sheet fluorescence microscopy), fluorescent probes, and material processing are paving the way for us to overcome these limitations. This allows for detailed visualization of sub-cellular structures within 3D tissues, providing valuable insights into critical cellular processes and their response to engineered environments.

We have also discussed the importance of balancing various factors like spatial and temporal resolution, signal-to-noise ratio, and photobleaching to achieve optimal image quality. This review serves as a comprehensive guide for researchers navigating the growing toolbox of imaging techniques available for studying 3D engineered tissues and organoids, ultimately leading to a deeper understanding of cell-cell interactions, disease mechanisms, and the development of improved in vitro models for drug discovery and therapy.

High-resolution sub-cellular imaging in tissue engineering has a bright future ahead of it. For the purpose of investigating larger and more complex 3D tissues, it will be important to continue developing improved microscopy techniques with even deeper tissue penetration and decreased phototoxicity. One of these emerging methods is Single-Molecule Orientation-Localization Microscopy that uses fluorogenic probes that specifically bind to the structure of interest; thus, it detects and analyzes signals from individual molecules to achieve higher resolution by capturing detailed orientation and positional data (Zhou et al., 2024). Additionally, Single-Molecule Orientation-Localization can use a radially and azimuthally polarized multi-view reflector microscope to enhance the precision of measuring 6D spatial and orientational coordinates (3D position and 3D orientation) of single molecules within engineered tissues (Zhang et al., 2023). Lattice Light Sheet Microscopy is another emerging optical technique which uses a rapidly moving thin plane of light sheet. Unlike light-sheet fluorescence microscopy, in which the light sheets are uniform, the light sheet here is structured into a lattice pattern (Daugird et al., 2024). Also, the investigation of fluorescent probes to improve luminosity, biocompatibility, and multi-color capabilities will enable the concurrent imaging of several cellular structures and functions using these optical methods. Lastly, in order to process the massive amounts of data produced by these high-resolution imaging techniques efficiently, the development of automated image analysis techniques will be necessary. Likewise, approaches to glean useful information from these massive data sets, including machine learning, will be crucial (Kalkunte et al., 2024).