Targeted nanocarriers for microglia-selective cytochalasin D delivery to modulate neuroinflammation after acute brain injuries

Targeted nanocarriers for microglia-selective cytochalasin D delivery to modulate neuroinflammation after acute brain injuries | Authorea try { document.documentElement.classList.add('js'); } catch (e) { } var _gaq = _gaq || []; _gaq.push(['_setAccount', 'G-8VDV14Y67G']); _gaq.push(['_trackPageview']); (function() { var ga = document.createElement('script'); ga.type = 'text/javascript'; ga.async = true; ga.src = ('https:' == document.location.protocol ? 'https://ssl' : 'http://www') + '.google-analytics.com/ga.js'; var s = document.getElementsByTagName('script')[0]; s.parentNode.insertBefore(ga, s); })(); Skip to main content Preprints Collections Wiley Open Research IET Open Research Ecological Society of Japan All Collections About About Authorea FAQs Contact Us Quick Search anywhere Search for preprint articles, keywords, etc. Search Search ADVANCED SEARCH SCROLL This is a preprint and has not been peer reviewed. Data may be preliminary. 18 November 2025 V1 Latest version Share on Targeted nanocarriers for microglia-selective cytochalasin D delivery to modulate neuroinflammation after acute brain injuries Authors : Sabri Elyes Mohamed Sahnoun , Rozina Noristani , Paolo Alimonti , Jonas Pes , Lara Gubeljak , Shaista Ilyas , Eren Arik , … Show All … , Alexandru Florea , Eva Miriam Buhl , Bernd Neumaier , Sanjay Mathur , Jörg B. Schulz , Felix Manuel Mottaghy , and Pardes Habib 0000-0002-5771-216X [email protected] Show Fewer Authors Info & Affiliations https://doi.org/10.22541/au.176349281.18872463/v1 343 views 163 downloads Contents Abstract Information & Authors Metrics & Citations View Options References Figures Tables Media Share Abstract Targeting microglia to modulate neuroinflammation after acute brain injuries (ABIs) is promising but limited by poor blood–brain barrier (BBB) penetration and systemic toxicity of candidate agents. We developed and characterized a tunable, folate-functionalized, polydopamine (PDA)-coated mesoporous silica nanoparticle (FA-NP) system for microglia-selective delivery and intracellular release of cytoskeleton inhibitor Cytochalasin D (CytoD). FA-NPs exhibited ideal physicochemical properties for BBB penetration and pH-sensitive PDA shells enabled intracellular retention and gradual degradation. Using radiolabeled and fluorescein-isothiocyanate-functionalized FA-NPs, SPECT/CT imaging, γ-counting, autoradiography, and immunohistochemistry confirmed BBB penetration. Activated microglia upregulated folate receptors (FOLR1/2) and internalized FA-NPs via FOLR-mediated, dynamin-dependent endocytosis. CytoD-loaded FA-NPs (FA-NP[CytoD]) significantly reduced microglial migration, phagocytosis, ROS production, and proinflammatory cytokines, outperforming molecular CytoD while exhibiting notably lower toxicity compared to both CytoD and unfunctionalized particles. In organotypic brain slice models of hypoxia-reoxygenation and traumatic injury, FA-NP[CytoD] reduced inflammation at 24 and 96 hours. Systemic administration enabled brain accumulation with clearance over time. Our modular theranostic platform combines microglial targeting, controlled drug release, and real-time central nervous system imaging, offering translational potential for ABIs. Targeted nanocarriers for microglia-selective cytochalasin D delivery to modulate neuroinflammation after acute brain injuries Sabri E. M. Sahnoun 1,2 , Rozina Noristani 3 , Paolo Alimonti 2,4 , Jonas Pes 5 , Lara Gubeljak 5 , Shaista Ilyas 6 , Eren Arik 5 , Alexandru Florea 1,11 , Eva M. Buhl 7 , Bernd Neumaier 8,9 , Sanjay Mathur 6 , Jörg B. Schulz 5,10 , Felix M. Mottaghy 1,11 , Pardes Habib 3,5 * 1 Department of Nuclear Medicine, University Hospital Aachen, RWTH Aachen University, 52074 Aachen, Germany 2 Department of Oncology, University of Oxford, Oxford OX3 7DQ, United Kingdom 3 Department of Neurosurgery and Stanford Stroke Center, Stanford University School of Medicine, Stanford, CA 94305, USA 4 Department of Neurosurgery, Brigham and Women’s Hospital, Harvard Medical School, Boston, MA 02120, USA 5 Department of Neurology, University Hospital Aachen, RWTH Aachen University, 52074 Aachen, Germany 6 Institute of Inorganic and Materials Chemistry, University of Cologne, 50939 Cologne, Germany 7 Electron Microscopy Facility, Institute of Pathology, University Hospital Aachen, RWTH Aachen University, 52074 Aachen, Germany 8 Institute of Radiochemistry and Experimental Molecular Imaging, University Hospital Cologne, University of Cologne, 50937 Cologne, Germany 9 Institute of Neuroscience and Medicine, Forschungszentrum Jülich GmbH, 52425 Jülich, Germany 10 JARA-BRAIN Institute of Molecular Neuroscience and Neuroimaging, Forschungszentrum Jülich GmbH and RWTH Aachen University, 52074 Aachen, Germany 11 Department of Radiology and Nuclear Medicine, Maastricht University Medical Center, 6202 Maastricht, The Netherlands *Correspondence: Pardes Habib, MD, PhD Department of Neurosurgery and Stanford Stroke Center Stanford University School of Medicine 1201 Welch Road, MSLS P352 Stanford, CA 94305-5487 [email protected] Running title: Targeted neurotheranostics for microglial modulation Abstract Targeting microglia to modulate neuroinflammation after acute brain injuries (ABIs) is promising but limited by poor blood–brain barrier (BBB) penetration and systemic toxicity of candidate agents. We developed and characterized a tunable, folate-functionalized, polydopamine (PDA)-coated mesoporous silica nanoparticle (FA-NP) system for microglia-selective delivery and intracellular release of cytoskeleton inhibitor Cytochalasin D (CytoD). FA-NPs exhibited ideal physicochemical properties for BBB penetration and pH-sensitive PDA shells enabled intracellular retention and gradual degradation. Using radiolabeled and fluorescein-isothiocyanate-functionalized FA-NPs, SPECT/CT imaging, γ-counting, autoradiography, and immunohistochemistry confirmed BBB penetration. Activated microglia upregulated folate receptors (FOLR1/2) and internalized FA-NPs via FOLR-mediated, dynamin-dependent endocytosis. CytoD-loaded FA-NPs (FA-NP[CytoD]) significantly reduced microglial migration, phagocytosis, ROS production, and proinflammatory cytokines, outperforming molecular CytoD while exhibiting notably lower toxicity compared to both CytoD and unfunctionalized particles. In organotypic brain slice models of hypoxia-reoxygenation and traumatic injury, FA-NP[CytoD] reduced inflammation at 24 and 96 hours. Systemic administration enabled brain accumulation with clearance over time. Our modular theranostic platform combines microglial targeting, controlled drug release, and real-time central nervous system imaging, offering translational potential for ABIs. Keywords Precision Medicine, Nanomedicine, Theranostics, Targeted Drug Delivery, Neurotherapeutics, Stroke, TBI 1. Introduction Neurological disorders are a major global health challenge, accounting annually for about 11 million deaths 1 . Their prevalence has risen steadily over the past three decades, now affecting nearly one in two individuals across lifespan 2 . Among these conditions, acute brain injuries (ABIs), such as stroke and traumatic brain injury (TBI), stand out as particularly devastating, becoming a leading cause of chronic disability in adults worldwide 3 . A major obstacle to therapeutic intervention in these conditions is the blood–brain barrier (BBB), which inhibits the entry of over 98 % of small molecules and nearly all macromolecular therapeutics to the brain parenchyma 4–6 . Consequently, the development of therapies for ABIs is constrained, emphasizing the need for strategies that enable targeted delivery to injured brain regions 7–9 . Microglia, the resident macrophage-like cells of the central nervous system (CNS), are the primary responders to CNS injury and have emerged as key targets in several neurological diseases 10,11 . Derived from yolk-sac progenitors, microglia populate the CNS early in development, maintaining themselves throughout life by self-renewal mechanisms. These distinct immune cells make up 10 % of the total cell population in the brain 12–14 . Under homeostatic conditions, microglia contribute to synaptic pruning, debris clearance, neurogenesis, myelination, and immune surveillance 14,15 . Following brain injury, microglia rapidly detect damage-associated molecular patterns released by damaged neurons and shift toward a reactive phenotype characterized by the secretion of pro-inflammatory cytokines, chemokines and reactive oxygen species (ROS) 16–18 . While microglial responses aim to restore homeostasis after CNS injury, excessive or prolonged activation can result in secondary neuronal damage and exacerbates neurodegeneration 13,19–22 . Consequently, selective modulation of microglial activity, without ablation, represents a promising therapeutic strategy for mitigating neuroinflammatory damage 23 . Cytochalasin D (CytoD), an actin polymerization inhibitor, disrupts microglial functions such as migration and phagocytosis, thereby attenuating their activation and inflammatory response 24–26 . However, the pronounced systemic toxicity and inability of CytoD to cross the BBB have precluded its clinical testing 27 . To overcome these limitations, biocompatible nanotechnology-based delivery systems offer promising potential for transporting drugs into the brain with both spatial and cellular precision. Among available platforms, mesoporous silica nanoparticles (mSiO 2 -NPs) enable a tunable architecture for drug loading and functionalization. Their versatile nature allows efficient encapsulation of therapeutic agents, while tailored surface modifications enhance circulation time, targeting capacity, and control over drug release 28–31 . However, uncoated silica particles have been associated with dose-dependent cytotoxicity, microglial stress and inflammatory responses, highlighting the need for biocompatible surface modifications to ensure safety 32–39 . Polydopamine (PDA) coatings on nanocarriers represent a robust strategy to improve nanoparticle biocompatibility, offering high serum stability, extensive drug loading capacity, and pH-sensitive release in acidic environments such as endolysosomal compartments 40–45 . In addition, PDA-coated NPs, and not bare silica NPs have been shown to cross the BBB and attenuate neuroinflammation in rodent models 46,47 . Upon activation, microglia upregulate folate receptors (FOLR 1/2), which makes folic acid (FA) an effective target ligand for selective drug delivery to these cells 48–50 . FA-functionalized PDA-coated NPs presents a promising theranostic (loaded drug + imaging moieties) approach to enhance selective uptake by reactive microglia while minimizing off-target effects 48–50 . However, despite the growing interest in PDA-coated NPs, our understanding of their cellular uptake dynamics, intracellular trafficking, subcellular biodistribution, and retention times in brain-resident cells remains incomplete. Limited understanding about the transport of PDA-coated NPs across the BBB, the mechanisms underlying their selective uptake by activated microglia, and the spatiotemporal regulation of drug release within the CNS hinders their clinical translation and rational design as PDA-modified nanotherapeutics for neurological applications. Here, we introduce a radiolabeled, PDA-coated, FA-functionalized mSiO 2 nanoparticle (FA-NPs) platform to address these gaps and enable targeted theranostic modulation of microglia in acute brain injury. Our system enables dual radiolabeling with Gallium-68 ( 68 Ga) for positron emission tomography combined with computed tomography (PET-CT) and Lutetium-177 ( 177 Lu), while encapsulating CytoD for localized immunomodulation. We chose the gamma- and beta-emitting radionuclide 177 Lu for its long half-life (6.7 d), allowing for prolonged intracellular tracking to investigate trafficking and biodistribution mechanisms. This was complemented by 68 Ga, a short-lived PET nuclide (67 min), for early-phase imaging. Given the delayed onset of radiation-induced damage, the short experimental duration, low CNS cell proliferation, and the minimal non-therapeutic dose [< 0.01 Bq/cell], potential 177 Lu-induced radiotoxic effects were unlikely to interfere with the mechanistic insights provided in this study. The modularly engineered and radiolabeled nanoparticles exhibit uniform morphology, high radiochemical purity, and excellent serum stability. FA-NPs are predominantly internalized through FOLR-mediated, dynamin-dependent endocytosis by microglia in the brain, with uptake further enhanced upon activation. Once inside endolysosomal compartments, acidic degradation of the PDA shell enables pH-responsive CytoD release, leading to suppression of microglial migration, phagocytosis, and inflammatory cytokine expression. In vivo single photon emission computed tomography (SPECT), post-mortem autoradiography (ARG), and immunohistochemistry (IHC) confirmed efficient BBB crossing, homogenous brain accumulation, and progressive clearance, demonstrating biocompatibility and translational potential. Using in vitro , in vivo, as well as hypoxia-reoxygenation and focal brain injury models, we elucidate the cellular mechanism of nanoparticle uptake, drug release, biodistribution, and microglial immunomodulation. This work introduces modularly architected FA-NP[CytoD] as a multifunctional nanotheranostic platform capable of microglia-directed modulation, real-time imaging, and controlled therapeutic delivery. By integrating receptor-mediated targeting, pH-sensitive payload release, and radiotracer-based biodistribution tracking, our system offers mechanistic insights into nano-bio interactions in the injured brain and introduces a strategy for precision neuroimmune modulation in CNS injury. 2. Results 2.1. Click- and DOTA-functionalized PDA-coated mSiO 2 -NPs display tunability, high radiochemical yield, purity, and stability in human sera Figure 1: Chemical and radiochemical functionalization, characterization, and stability of modified mSiO 2 nanoparticles. (A) Schematic illustration of template-assisted sol-gel synthesis of mSiO 2 -NPs, surface coating with polydopamine (PDA) to generate PDA-NPs, carbodiimide coupling reaction for the conjugation of folic acid on PDA-capped mSiO 2 -NPs (FA-NP), followed by chelator (DOTA) attachment, CytoD loading and radiolabeling with the theranostic pair 68 Ga and 177 Lu. (B) FT-IR spectra of mSiO 2 (black), PDA-NP (red), FA-NP (blue), and pure FA (green) modified from Ilyas et al . 51 . The surface charges (zeta potential) for the bare and functionalized mSiO 2 -NPs in water at pH 7 in triplicates are shown in the table. (C-E) The hydrodynamic diameter [nm] and polydispersity index (PDI) were assessed using nanoparticle tracking analysis (NTA). Representative TEM images of mSiO 2 -NPs and their functionalized derivatives are presented. The radiolabeling efficiency of 68 Ga- (F) and 177 Lu- (G) labeled NPs were assessed utilizing radio-thin layer chromatography (radio-TLC). (H) Evaluation of the lipophilicity/partition coefficient (P) and the calculated logP value of NPs were measured using a gamma counter (γ-Counter). (I) Radio-TLC analysis of the concentration-dependent radiochemical yield (RCY) of 68 Ga- and 177 Lu-labeled NPs ([ 68 Ga]Ga-DOTA-NP = 68 Ga-DOTA-NP; [ 177 Lu]Lu-DOTA-NP = 177 Lu-DOTA-NP). To determine the serum stability, 68 Ga- (J) and 177 Lu- (K) labeled NPs were measured in human sera and PBS control in a time-dependent manner. Data are shown as means ± SD of 3 independent experiments with 3 technical replicates. To engineer a tunable nanostructured platform for theranostic microglial modulation, we designed a modularly architected nanoparticle system integrating targeted delivery, pH-sensitive drug release, and radionuclide-based imaging capabilities. The multi-functionalization, characterization, and stability of mSiO 2 -NPs were successfully assured through a series of chemical modifications and a subsequent radiolabeling procedure. Bare mSiO 2 nanoparticles were synthesized via a template-assisted sol-gel process (TEOS), followed by polydopamine (PDA) coating to form PDA-NPs. Subsequently the NPs were conjugated with folic acid (FA) using carbodiimide chemistry to create PDA-capped and FA functionalized mSiO 2 -NPs (FA-NPs) (Fig. 1A). The conjugation was confirmed through Fourier-transform infrared spectroscopy (FTIR) as previously shown 51 , where the appearance of a peak at 1696 cm⁻¹, corresponding to the C=O vibration, indicated remaining carboxy groups post-FA conjugation, while a peak at 1604 cm -1 confirmed the presence of amide bonds formed during the coupling reaction (Fig. 1B). Zeta potential measurements further confirmed the successful functionalization of the nanoparticles. The surface charges shifted from -28.5 mV (mSiO 2 -NP) to -41.2 mV (PDA-NP) and -37.5 mV (FA-NP) (Fig. 1B). Morphological characterization using transmission electron microscopy (TEM) revealed a spherical mesoporous structure with a mean diameter of approximately 110 nm for mSiO 2 -NPs. PDA shell measurements revealed an average thickness of 29 nm with uniformly distributed pores (~4 nm in diameter). Furthermore, nanoparticle tracking analysis (NTA) detected hydrodynamic diameters in the range of 220-260 nm with polydispersity indices (PDI) below 0.15, indicating uniform size distribution across all three NP formulations (Fig. 1C-E). Radiolabeling was achieved by covalently attaching the radiometal chelator DOTA by conjugation of p-SCN-Bn-DOTA (S-2-(4-isothiocyanatobenzyl)-1,4,7,10-tetraazacyclododecane tetraacetic acid) with the NP surface, which allowed chelation of both the PET-imaging radionuclide 68 Ga (t ½ = 68 min) and the therapeutic radionuclide 177 Lu (t ½ = 6.7 d). Radio-TLC displayed radiolabeling conversions > 98 % for both isotopes. The absence of a [ 68 Ga]Ga-DOTA or [ 177 Lu]Lu-DOTA peak at around 50 mm position of the TLC indicates that DOTA remained stably bound to the nanoparticle; such a peak would be expected if DOTA had dissociated (Fig. 1F&G). To exclude the possibility that the choice of radionuclide influences the nanoparticle uptake rate, we compared competitive uptake of [ 68 Ga]Ga-DOTA-PDA-NP and [ 177 Lu]Lu-DOTA-FA-NP (1:1) versus [ 177 Lu]Lu-DOTA-PDA-NP and [ 68 Ga]Ga-DOTA-FA-NP (1:1) in all three neuronal cells. The type of radionuclide appeared to have no impact on uptake rates in microglia, astrocytes, and neurons (Figure S1 A-D, Supporting Information). Additionally, partition coefficient (logP) measurements using a γ-Counter revealed a calculated value of -3.31 ± 0.09, indicating the hydrophilic nature of the radiolabeled NPs (Fig. 1H). Optimal radiolabeling was observed at NP concentrations of ≥ 0.5 mg/mL for 68 Ga and ≥ 0.7 mg/mL for 177 Lu, with carrier concentrations reaching up to 0.15 ng (Lu)/µL (Fig. 1I). The stability of the radiolabeled NPs was further confirmed in human sera at 37 °C. Radio-TLC analysis demonstrated that the 68 Ga-labeled NPs remained stable (RCY > 97 %) for over 3 h, while the 177 Lu-labeled NPs maintained stability (RCY > 95 %) for up to 120 h (Fig. 1J&K). In summary, we modularly engineered mSiO 2 -NPs functionalized with PDA and FA, and stably radiolabeled with 68 Ga and 177 Lu, achieving over 98 % labeling yield. Characterization confirmed uniform size, tunable functionalization, and high stability in human sera. Competitive uptake studies using two different radionuclides show no effect of the radionuclide type on uptake in brain cells, supporting their combined diagnostic and therapeutic potential. 2.2. Activated microglia upregulate folate receptors and exhibit increased uptake of FA-functionalized NPs Figure 2: Impact of NP surface functionalization on cellular uptake, intracellular localization, and distribution in neuronal cells. Cellular uptake rates using immunocytochemistry (ICC), transmission electron microscopy (TEM) and γ-Counter in (A) microglia (HMC-3), (B) astrocytes (U373), and (C) neurons (SY5Y). ICC co-staining of fluorophore-labeled FITC-FA-NP (green), DAPI (blue), and cell-specific markers IBA1 for microglia, GFAP for astrocytes, and NeuN for neurons (red) (A-C top panel) . Representative TEM images showed accumulated FA-NPs in the cytoplasm of all three cell lines captured after 1 h of NP incubation (A-C middle panel) . Decay-corrected quantitative cellular uptake rates of PDA-NP and FA-NP (control or 3 h LPS [1µg/mL] treatment) were measured by γ-Counter using 177 Lu-labeled NPs [< 0.01 Bq/cell] (A-C bottom panel) . Data are shown as means ± SD of 9 independent experiments with 2 technical replicates . A two-way ANOVA followed by multiple comparison tests and Tukey’s post-hoc test was performed. ***p < 0.001; *p < 0.05. Representative TEM images of mSiO 2 -NPs (D) , PDA-NPs (E) , and FA-NPs (F) after 1 h of NP incubation in HMC-3 microglial cells. ICC analysis of HMC-3 microglial cells with FITC-FA-NPs, DAPI, FOLR1 (G) , and FOLR2 (H) antibodies. Western blot analysis of FOLR1 (I) and FOLR2 (J) with actin serving as loading control. Data are shown as means ± SD of 3 independent experiments with 3 technical replicates. A Kruskal-Wallis test followed by Dunn’s multiple comparisons test was performed . *p < 0.05. To evaluate the potential of FA-functionalized, PDA-coated mSiO 2 -NPs (FA-NPs) for targeted drug delivery in neuronal cells, we visualized and quantified the intracellular uptake using ICC, TEM and γ-Counter (Fig. 2A-C). To visualize NP internalization, FITC-labeled FA-NPs (FITC-FA-NPs) were administered to microglia, astrocytes, and neurons for 1 h. ICC demonstrated that FITC-FA-NPs co-localized with cell-specific markers IBA1 (microglia), GFAP (astrocytes), and NeuN (neurons), indicating visual internalization across all three cell types (Fig. 2A-C, top panel). TEM analysis displayed internalized FA-NPs in all three cell types, predominantly localized within vesicular structures in cytoplasm and perinuclear regions. Notably, microglia exhibited a more dispersed vesicular distribution, with FA-NPs present in multiple endolysosomal compartments throughout the cytoplasm (Fig. 2A-C, middle panel). To quantify uptake rates of 177 Lu-labeled FA-NPs and PDA-NPs, we utilized a γ-Counter after 1 h of NP incubation. To mimic an inflammatory/activated state of neuronal cells, we subjected all three cell types to lipopolysaccharide (LPS) treatment [1 µg/mL] or vehicle for a total of 3 h (Fig. 2A-C, bottom panel). To investigate the effect of surface functionalization, we compared the uptake rates of unfunctionalized, PDA-coated mSiO 2 -NPs (control: PDA-FP) and FA-functionalized, PDA-coated mSiO 2 -NPs (FA-NPs) across all three cell types. A significant increase in uptake was observed exclusively in microglial cells following FA functionalization (p < 0.001). Microglia also displayed the highest uptake rates of FA-NPs (~40 %), followed by astrocytes (~20 %) and neurons (~11 %). Notably, LPS treatment further increased microglial uptake of FA-NPs (~48 %, p < 0.05), whereas LPS had no effect on FA-NP uptake in astrocytes and neurons (Fig. 2A-C, bottom panel). To further investigate the enhanced uptake of FA-functionalized NPs in microglial cells, we performed TEM imaging to visualize the internalization of bare mSiO 2 -NPs, PDA-NPs, and FA-NPs, and assessed folate receptor expression using ICC and Western blot analysis (Fig. 2D-J). Consistent with our previous findings, FA functionalization led to markedly increased intracellular accumulation compared to bare mSiO 2 -NPs and PDA-NPs (Fig. 2D–E). Microglial cells expressed both folate receptors (FOLR1/FOLR2), and Western blot quantification revealed a significant upregulation of both receptors upon LPS stimulation (p < 0.05). In conclusion, FA-NPs are preferentially internalized by microglia, and this uptake is further amplified by microglial activation, which is accompanied by upregulation of folate receptors FOLR1 and FOLR2. This activation-dependent enhancement is specific to microglia and not observed in astrocytes or neurons. The vesicular localization of internalized particles suggests uptake through an endolysosomal pathway, potentially involving folate receptor–mediated endocytosis. 2.3. Microglia rapidly internalize FA-NPs via FOLR- and dynamin-dependent endocytosis Figure 3: Uptake mechanisms of FA-NPs in human mixed neuronal cell cultures and murine organotypic brain slice cultures. (A) Representative ICC images of FITC-FA-NPs in mixed human neuronal cell cultures 5:3:2 (microglia: astrocytes: neurons) with cell-specific markers for HMC-3 (IBA1), U373 (GFAP), and SY5Y (NeuN) after 1 h of FA-NP incubation. (B) Representative coronal murine brain slices (top panel) are 350 µm thick and preserve intact brain architecture. Immunohistochemical (IHC) analysis (bottom panel) of FITC-FA-NP uptake in adult murine organotypic brain slice cultures (12 µm sections) following 1 min, 30 min, and 60 min of FA-NP incubation. (C) Schematic overview of pharmacological inhibitors used to dissect uptake mechanisms. (D-F) Decay-corrected γ-Counter analysis of cellular 177 Lu-FA-NP [< 0.01 Bq/cell] uptake following 1 h of NP incubation in HMC-3, U373, and SY5Y cells treated for 30 min with free FA [1 mM], CytoD [10 µM], Rottlerin [6 µM], or Dynasore [10 µM]. Concentration-dependent quantitative 177 Lu-FA-NP [< 0.01 Bq/cell] uptake in microglia was measured by γ-Counter after a FA-NP incubation time of 1 h using different molar concentrations of Rottlerin (G) and Dynasore (H) (treated 30 min prior to NP incubation). (I) Combined inhibition studies in microglia using co-treatment of free FA [1 mM] in combination with Rottlerin [6 µM] or Dynasore [10 µM] for 30 min prior to 177 Lu-FA-NP [< 0.01 Bq/cell] incubation for 1 h. Data are shown as means ± SD of 3-6 independent experiments with 2-3 technical replicates . A one-way ANOVA followed by Dunnett’s multiple comparisons test was performed. ***p < 0.001; **p < 0.01; *p < 0.05. To investigate cellular uptake mechanisms of FA-functionalized NPs, we first assessed their internalization in mixed human neuronal cell cultures composed of microglia, astrocytes, and neurons at a physiologically relevant ratio of 5:3:2. ICC analysis revealed robust uptake of FITC-FA-NPs across all three cell types, with microglia exhibiting markedly higher uptake rates than astrocytes and neurons (Fig. 3A). We next utilized murine organotypic brain slice cultures to preserve the three-dimensional cytoarchitecture of the brain, to evaluate spatiotemporal cellular uptake of FITC-FA-NPs. Following 1, 30, and 60 minutes of incubation, FITC-FA-NP uptake increased over time in all cell types. Notably, strong co-localization of FITC-FA-NPs with the microglial marker IBA1 was observed as early as 1 minute post-incubation, suggesting rapid uptake kinetics in microglia (Fig. 3B). Uptake by astrocytes and neurons was also detectable but substantially less prominent. To dissect the mechanisms underlying FA-NP internalization, we pretreated neuronal cells with pharmacological inhibitors targeting distinct endocytic pathways: free folic acid (FA) to competitively block folate receptors (FOLR), Cytochalasin D (CytoD) to disrupt actin polymerization and phagocytosis, Rottlerin to inhibit macropinocytosis and non-clathrin pathways, and Dynasore to block dynamin-dependent endocytosis (Fig. 3C). We tested and utilized non-toxic molar concentrations of the inhibitors as previously described 52–54 (Fig S2 A&B, Supporting Information). Administration of CytoD [10 µM] had no significant impact on 177 Lu-FA-NP uptake in any of the tested cell types, indicating that phagocytosis is not a major uptake route (Fig. 3D-F, Figure S2 A&B, Supporting Information). In contrast, treatment with Rottlerin [6 µM] and Dynasore [10 µM] led to a significant reduction in 177 Lu-FA-NP uptake in all cell types, with the strongest effect observed in microglia (p < 0.001). Dose-dependent inhibition by both Rottlerin and Dynasore further confirmed their impact on microglial uptake (Fig. 3G&H). While treatment with free FA [1 mM] did not significantly reduce 177 Lu-FA-NP uptake in astrocytes and neurons, a significant decrease was observed in microglia (p < 0.001, Fig. 3D-F) indicating a FOLR-mediated endocytic uptake. We next pre-incubated HMC-3 microglial cells with free FA [1 mM] + Rottlerin [6 µM] or free FA [1 mM] + Dynasore [10 µM] for 30 min prior to 177 Lu-FA-NP administration to assess the combined inhibition of FOLR-mediated endocytosis. The combination of free FA and Rottlerin did not result in significant difference compared to treatment with rottlerin alone. In contrast, co-treatment of free FA + Dynasore revealed a significant reduction of 177 Lu-FA-NP uptake in microglia (p < 0.001) (Fig. 3I). Collectively, these data indicate that FA-functionalized NPs are rapidly and predominantly internalized by microglia via dynamin- and FOLR-dependent endocytosis, while astrocytes and neurons display lower uptake. These findings underscore microglia as the principal cellular target of FA-NPs in the brain, offering insight into cell-specific delivery strategies for neurotherapeutics. 2.4. FA-NPs are rapidly internalized and retained in microglia for up to 96 hours Figure 4: Time-dependent uptake and intracellular retention time of FA-NPs in microglia. (A ) Schematic overview of the experimental workflow for time-resolved uptake and retention assays using FITC- and 177 Lu-labeled FA-NPs. (B) Time-dependent uptake of FITC-FA-NPs (green) in microglia using ICC staining (DAPI (blue) and IBA1 (red)). (C) Diagram of the γ-Counter setup used to measure 177 Lu-FA-NP accumulation in cell lysates and supernatants. (D) Decay-corrected quantitative γ-Counter uptake rates of 177 Lu-labeled FA-NPs [< 0.01 Bq/cell] up to 3 h of NP incubation. (E) Time-dependent retention measurements using ICC analysis with FITC-FA-NPs (green), DAPI (blue), and Lysotracker (red) up to 96 h of intracellular retention. (F) Decay-corrected quantitative γ-Counter measurements with 177 Lu-labeled FA-NPs [< 0.01 Bq/cell] up to 96 h of intracellular retention. Data are shown as means ± SD of 3 independent experiments with 3 technical replicates . A one-way ANOVA followed by Dunnett’s multiple comparisons test was performed. ***p < 0.001; *p < 0.05. To investigate the kinetics of microglial interaction with FA-NPs, we performed time-resolved uptake and retention studies using both 177 Lu-labeled and FITC-functionalized FA-NPs (Fig. 4A). ICC revealed a detectable internalization of FA-NPs (green) as early as 1 minute post-incubation, with progressive cytoplasmic accumulation peaking between 30 and 60 minutes (Fig. 4B). Co-localization (yellow) with the microglial marker IBA1 confirmed cell-specific uptake, while DAPI-staining delineated nuclear morphology. These findings were corroborated by quantitative γ-Counter analysis (Fig. 4C), which demonstrated rapid and significant NP uptake within the first minute (approx. 5%), increasing steadily over time (p < 0.001). Uptake saturated at approximately 40 % after 60 minutes, with no further increase at 120 or 180 minutes, indicating that internalization reached a plateau within one hour (Fig. 4D). To assess the intracellular retention time, we tracked NP localization using ICC with Lysotracker, revealing co-localized FITC-FA-NP signal within lysosomal compartments up to 96 hours post-incubation (Fig. 4E). This was supported by γ-Counter measurements, which showed a gradual decrease in cell-associated radioactivity over time, accompanied by a corresponding increase in signal within the supernatant, consistent with NP efflux or release (Fig. 4F). Together, these data indicate that FA-NPs are rapidly and efficiently internalized by microglial cells, reaching maximum uptake within 60 minutes. Although intracellular levels decline over 96 hours, a substantial fraction of FA-NPs is retained long-term, suggesting endolysosomal sequestration with partial release into the extracellular environment over time. 2.5. Intracellular pH triggers PDA shell degradation and cargo release from FA-NPs Figure 5: pH-dependent PDA-shell degradation and intracellular drug release of FA-NPs in microglia. (A) Representative TEM images of FA-NPs internalized in HMC-3 microglial cells, showing influence on PDA coating in early and late endosomes (E) (red arrow = decreased PDA shell; black arrow = mostly intact PDA shell). PDA mean thickness r (mean of four different shell locations) (B) and PDA mean area (C) of FA-NPs incubated in media (48 h at pH 7.4 or 5.0, respectively) and after an intracellular retention time of 48 h or 96 h in HMC-3 microglial cells were measured using ImageJ. (D) Time-dependent cargo release in ICC staining of DAPI-loaded FITC-FA-NPs in microglia, co-stained with a Lysotracker. Bars represent means ± SD. 50-100 FA-NPs per condition were analyzed. A two-tailed t-test followed by Welch’s test was performed. ***p < 0.001. To analyze the intracellular stability and release behavior of polydopamine (PDA)-coated folic acid-functionalized nanoparticles (FA-NPs), we investigated PDA shell degradation under acidic conditions and monitored NP-mediated delivery of DAPI cargo in microglial cells. Transmission electron microscopy (TEM) revealed that PDA shells remained largely intact in early endosomal compartments, while degradation was evident in larger vesicles corresponding to late endosomes (Fig. 5A). Morphological changes included thinning of the PDA layer (black vs. red arrows) and vesicle fusion events (marked with *), suggestive of dynamic endolysosomal processing (Fig. 5A). Quantification of PDA-shell thickness and surface under different pH conditions confirmed that acidic environments promote the degradation process. FA-NPs subjected to an experimental pH value of 5.0 displayed a significant degradation of the PDA-shell thickness and area compared to those maintained at physiological pH 7.4 (p < 0.001) (Fig. 5B&C, Figure S3 A&B, Supporting Information). Similarly, intracellular PDA-shell degradation was evident by 48 h post-uptake in microglial cells, with further reduction at 96 h (p < 0.001), indicating progressive degradation over time (Supplementary Fig. 3C–D). Notably, the PDA shells were not completely degraded at either time point, suggesting that a residual shell structure persists even after prolonged intracellular retentions times. To assess functional cargo release capacity of FA-NPs, we encapsulated DAPI within FITC-labeled FA-NPs and monitored intracellular localization using immunocytochemistry. A time-dependent increase in cytosolic FITC signal was observed alongside progressive nuclear DAPI staining, starting as early as 5 minutes post-incubation and culminating in complete nuclear labeling by 60 minutes (Fig. 5D). Co-staining with Lysotracker revealed partial overlap, supporting endolysosomal involvement in cargo release. These data demonstrate that PDA-coated FA-NPs exhibit acid-sensitive degradation and support sustained intracellular release of cargo. Importantly, the observation that cargo release occurs despite incomplete PDA shell degradation supports the notion that full shell degradation is not required for effective delivery. This may point to a mechanism involving partial endosomal escape or localized shell disruption. Together, these findings highlight the utility of FA-NPs as a pH-responsive nanocarrier system capable of overcoming endolysosomal barriers for targeted intracellular delivery. 2.6. FA-NP encapsulated CytoD attenuate toxicity and selectively modulate microglial inflammatory response, migration and phagocytosis Figure 6: Impact of PDA coating, FA functionalization, and CytoD encapsulation on toxicity, ROS production, inflammatory response, phagocytosis, and migration of human microglial cells. Cellular toxicity and inflammatory response were measured both at 24 h and 96 h post-incubation (incubation time 1 h) for bare mSiO 2 -NPs, PDA-NPs, FA-NPs, FA-NPs-encapsulated CytoD (FA-NP[CytoD]), molecular CytoD [5 µM] and the corresponding NaCl control. (A) LDH release, (B) cell viability using cell counter, (C) metabolic activity, (D) ROS production and (E) impact on inflammatory cytokines and pro-apoptotic genes. Data are shown as means ± SD of 3-4 independent experiments with 3 technical replicates. A Kruskal-Wallis test followed by Dunn’s multiple comparisons test was performed . ***p < 0.001; **p < 0.01; *p < 0.05. Migration ability was assessed using a zone exclusion assay. (F) Representative images of microglial HMC-3 migration over a period of 48 h after stimulation. (G) Quantification of HMC-3 migration treated either with NaCl (control), FA-NP [6.25 µg/mL], FA-NP[CytoD] [6.25 µg/mL] and molecular CytoD [5 µM] up to 48 h. Data are presented as means ± SEM of 6 individual experiments with 2 technical replicates in each experiment. (H) Representative images of phagocytic activity are shown. Phagocytic activity of microglial HMC-3 cells using pHrodo TM Red E. coli Bioparticles TM after 1 h of treatmet with either NaCl (control), FA-NP [6.25 µg/mL], FA-NP[CytoD] [6.25 µg/mL], molecular CytoD [5 µM] and LPS (positive control, 1 µg/mL) was assessed at 24 and 48 h. (I) Quantification of phagocytic activity. pHrodo TM Red E. coli Bioparticles TM positive cells per 1,000 counted cells were quantified in each experiment. Data are shown as means ± SD of 4 independent experiments with 3 technical replicates. A Kruskal-Wallis test followed by Dunn’s multiple comparisons test was performed . ***p < 0.001; **p < 0.01; *p < 0.05. Following our observation of efficient FA-NPs uptake by microglial cells, we investigated the downstream impact of NP exposure and CytoD delivery on microglial toxicity, oxidative stress, inflammatory response, and key microglial functions such as migration and phagocytosis. An exposure to bare mSiO 2 -NPs, PDA-NPs, and FA-NPs for 1 h followed by washout revealed no acute toxicity at 24 h, as assessed by LDH release or cell viability after 24 hours (Fig. 6A&B). However, at 96 h, the bare mSiO 2 -NPs particles exhibited significantly increased LDH release and reduced microglial viability. In contrast, PDA and FA functionalization attenuated these effects, demonstrating a protective impact of surface modification. Free molecular CytoD [5 µM] induced significant cell death at both 24 h and 96 h post-incubation, while FA-NP-encapsulated CytoD (FA-NP[CytoD]) significantly reduced LDH release and cell death (Fig. 6A&B). Notably, the bare mSiO 2 -NPs revealed decreased metabolic activity and increased ROS production in microglial cells at both 24 and 96 h (Fig. 6C&D). Interestingly, NPs with PDA coating and FA functionalization did not exhibit a significant reduction in metabolic activity compared to NaCl controls at both time points, indicating a protective effect of the functionalized NPs (Fig. 6C). Furthermore, functionalization with PDA and FA substantially reduced ROS production, suggesting a reduction in oxidative stress (Fig. 6D). In addition to increased toxicity, bare mSiO 2 -NPs also elevated the expression of pro-inflammatory cytokines (IL-6, IL-1β, TNF, CCL2, and CCL5) and pro-apoptotic genes (BAX, CASP3, CASP8, and CASP9) at both 24 h and 96 h (Fig. 6E). PDA-coated and FA-functionalized NPs suppressed both inflammatory and apoptotic gene expression, highlighting the anti-inflammatory profile of functionalized particles and their capacity to reduce activation-associated toxicity (Fig. 6E). Given the reduced toxicity and inflammatory response observed with FA-NPs-encapsulated CytoD, we next assessed the impact of these NPs on key microglial functions such as migration and phagocytosis. Microglial migration was evaluated using a zone exclusion assay with time-lapse imaging over 48 h (Fig. 6F&G). FA-NPs significantly enhanced microglial migration compared to the NaCl control. However, the administration of molecular CytoD [5 µM] effectively inhibited migration. Encapsulation of CytoD within FA-NPs also led to a notable reduction in migration compared to FA-NPs alone, demonstrating that FA-NP[CytoD] effectively inhibit microglial mobility (Fig. 6F&G). With increasing concentrations of FA-NP-encapsulated CytoD [0–12.5 µg/mL], microglial cell migration decreased over a 48 h observation period (Figure S4 A, Supporting Information). The administration of the endocytosis inhibitor Dynasore counteracted the inhibitory effect of CytoD on cell migration, further suggesting the endocytic uptake of the nanoparticles (Figure S4 5B&C, Supporting Information). To investigate phagocytic activity, microglial cells were incubated with pH-dependent coupled beads (pHrodo™) for 24 h and 48 h (Fig. 6H&I). CytoD significantly reduced phagocytosis at both time points, and NP-encapsulation of CytoD within FA-NPs similarly diminished microglial phagocytosis. In contrast, LPS [1 µg/mL] used as a positive control elicited the highest levels of phagocytosis, as expected (Fig. 6H&I). In Summary, PDA coating and FA functionalization of NPs effectively reduce microglial toxicity, ROS production, and the inflammatory response. Furthermore, encapsulation of CytoD within FA-NPs preserves its functional ability to inhibit microglial migration and phagocytosis, while mitigating its cytotoxic effects. This suggests that functionalized FA-NPs may serve as a promising platform for modulating microglial activity in neuroinflammatory conditions. 2.7. FA-NP-encapsulated CytoD mitigate neuroinflammation following hypoxia-reoxygenation or focal brain injury Figure 7: Impact of FA-NP-encapsulated CytoD on proinflammatory cytokine transcript levels in organotypic adult murine brain slice cultures after hypoxia-reoxygenation or focal thermocoagulation injury. (A) Protocol of hypoxia-reoxygenation injury in organotypic adult murine brain slice cultures (350 µm sections). (B) Schematic illustration of hypoxia-chamber with (C) hypoxia conditions. (D) Time-dependent (24 h or 96 h post-NP incubation) γ-Counter uptake rates of 177 Lu-labeled FA-NPs [< 0.01 Bq/cell]) in organotypic adult murine brain slice cultures are presented (N = normoxia; H = hypoxia). mRNA levels of IL-1β (E) , IL-6 (F) , and TNF (G) were assessed 24 h or 96 h post-hypoxia/normoxia after NP incubation. Brain slices were incubated with FA-NP or FA-NP[CytoD] [6.25 µg/mL] for 1 h in both injury models. (H) Protocol of focal thermocoagulation injury in organotypic adult murine brain slice cultures (350 µm / slice). (I) A schematic illustration of cortical/subcortical focal thermocoagulation injury. (J) Representative picture and autoradiography (ARG) of a coronal organotypic adult murine brain slice (350 µm sections) after focal thermocoagulation and incubation with 177 Lu-labeled FA-NPs [< 0.01 Bq/cell]. (K) Time-dependent (24 h or 96 h post-NP incubation of 1 h) γ-Counter uptake rates of 177 Lu-labeled FA-NPs [< 0.01 Bq/cell] in ipsi- and contra-lesional brain hemispheres. mRNA levels of IL-1β (L) , IL-6 (M), and TNF (N) were assessed 24 h or 96 h after focal brain injury. Intergroup differences were tested by two-way ANOVA followed by Tukey´s post-hoc test (multiple comparisons). Bars represent means ± SD of 4-6 individual experiments with 3 technical replicates in each experiment. ***p < 0.001; **p < 0.01; *p < 0.05. Acute hypoxic/ischemic and traumatic focal brain injuries represent the most common causes of neurological morbidity, characterized by rapid neuroinflammatory responses and secondary tissue damage. Given the observed reduction of inflammatory and pro-apoptotic markers in microglial cells upon FA-NP-encapsulated CytoD treatment, we next evaluated its therapeutic potential in two ex vivo models of acute neurological injury: hypoxia-reoxygenation and focal thermocoagulation in adult murine brain slice cultures. In the hypoxia-reoxygenation model, organotypic adult murine brain slices (350 µm) were subjected to 30 min of hypoxia followed by reoxygenation for up to 96 h, with FA-NPs incubated for 1 h during the hypoxic period (Fig. 7A–C). Uptake rates of 177 Lu-labeled FA-NPs, measured via γ-counting, showed a significant increase in the hypoxic group at 24 h post-incubation (p < 0.001), while no significant difference in NP uptake was observed after 96 h in comparison to the normoxia group (Fig. 7D). As expected, hypoxia-reoxygenation injury led to elevated mRNA expression of pro-inflammatory cytokines IL-1β, IL-6, and TNF. While FA-NPs alone did not alter cytokine expression, treatment with FA-NP[CytoD] significantly suppressed all three cytokines at both 24 h and 96 h post-injury (Fig. 7E–G). To assess performance in the focal brain injury model, we employed a focal thermocoagulation injury model targeted to the motor cortex and adjacent subcortical structures, including the corpus callosum (Fig. 7H–I). Autoradiography showed lower FA-NP uptake in the ipsilesional hemisphere relative to the contralateral side, consistent with compromised local tissue integrity (Fig. 7J). γ-Counter quantification confirmed significantly greater FA-NP uptake in the contralateral hemisphere at 24 h (p < 0.01), which normalized by 96 h (Fig. 7K). Inflammatory gene expression was then assessed in ipsi- and contralateral hemispheres. In the ipsilesional tissue, FA-NP[CytoD] significantly reduced IL-1β, IL-6, and TNF mRNA expression at both 24 h and 96 h, compared to both NaCl and FA-NP controls (p < 0.001) (Fig. 7L–N). Similar anti-inflammatory effects were observed in the contralateral hemisphere, with significant reductions in IL-1β (24 h), IL-6 (96 h), and TNF (both time points). Collectively, these findings demonstrate that FA-NP-encapsulated CytoD effectively suppresses inflammatory cytokine expression in models of ischemic-like and traumatic brain injury and support its potential as a promising nanotherapeutic candidate to be further evaluated in in vivo models of acute brain injury. 2.8. Systemic administration of FA-NPs enables brain delivery and cellular uptake with partial clearance by 24 hours Figure 8: Time-dependent in vivo biodistribution, quantification, and retention of FA-NPs in murine brains. (A) Schematic experimental procedure: 177 Lu-labeled and FITC-functionalized FA-NPs (30 MBq) were intravenously injected into mice. SPECT/CT imaging was performed at 4 h (n = 3) or 24 h (n = 3) post-injection (p.i.), followed by post-mortem (p.m.) ARG, γ-Counter quantification, and IHC analysis at 24 h and 120 h. (B) SPECT/CT images at 4 h or 24 h p.i. show the distribution of 177 Lu-labeled FA-NPs in the brain. (C) ARG of 2 mm coronal brain sections at 24 h or 120 h post-injection. (D) Decay-corrected quantitative γ-Counter analysis, represented as counts per minute (CPM) for individual brain sections, shows the FA-NP retention over time at 24h or 120 h. A one-way ANOVA followed by multiple comparison tests and Tukey’s post-hoc test was performed. ***p < 0.001. IHC analysis of brain sections at 24 h (E) or 120 h (F) p.i. demonstrates the localization of FITC-FA-NPs within the brain using markers for microglia (IBA1), astrocytes (GFAP), and neurons (NeuN). To assess the in vivo biodistribution, BBB penetration capacity, and clearance profile of FA-NPs, mice were intravenously injected with 177 Lu-labeled (30 MBq) FITC-FA-NPs, followed by SPECT/CT imaging at 4 h and 24 h post-injection (Fig. 8A). SPECT/CT revealed an increased FA-NP accumulation in the brain after 4 h, consistent with successful BBB penetration followed by a decline in signal intensity at 24 h, suggesting partial clearance or degradation of the nanoparticles over time (Fig. 8B). Post-mortem autoradiography (ARG) of serial coronal brain sections supported the imaging data, with widespread radiotracer accumulation observed at 24 h, and notably reduced signal at 120 h (Fig. 8C). Quantitative γ-Counter analysis confirmed this decline in nanoparticle retention, showing significantly higher counts per minute (CPM) at 24 h compared to 120 h (p < 0.001; Fig. 8D). To determine cellular distribution, immunohistochemistry (IHC) was performed using markers for microglia (IBA1), astrocytes (GFAP), and neurons (NeuN). At 24 h p.i., FITC-FA-NPs were clearly detectable in all three CNS cell types (Fig. 8E). By 120 h p.i., FITC signal was visibly diminished across all cell types, consistent with the SPECT/CT, ARG, and γ-Counter data (Fig. 8F). Together, these results confirm that FA-NPs can efficiently cross the BBB and reach key neural cell populations in vivo as early as 4 h post-injection. The signal decline after 120 h suggests clearance or degradation, supporting the potential for transient CNS delivery without long-term accumulation — an important feature for translational therapeutic applications. 3. Discussion Modulating prolonged or dysregulated microglial activation after acute brain injuries, including stroke, trauma, and hypoxic insults, holds great promise for mitigating secondary damage. However, systemic delivery of immunomodulatory drugs to the brain remains severely limited by the BBB, which prevents most therapeutics from accessing parenchymal targets. To address this challenge, we developed a multifunctional, tunable PDA-coated, FA-functionalized mSiO 2 -NP platform, designed for efficient BBB penetration, microglial targeting, controlled intracellular drug release, and imaging-guided tracking. mSiO 2 -NPs possess unique structural and physicochemical properties that confer chemical stability, tunable surface functionality, and excellent biocompatibility, establishing them as promising carriers for the targeted delivery of theranostic payloads 55–57 . Their mesoporous architecture facilitates drug encapsulation while reducing hemolytic activity compared to nonporous silica NPs, attributed to reduced silanol density 58,59 (Fig. 1A). Key parameters such as particle size and surface chemistry critically influence NP behavior in the organism 60 . While Napierska et al ., reported that mSiO 2 -NPs larger than 100 nm exhibit minimal endothelial toxicity, particles smaller than 50 nm can induce significant necrotic cell death 61 . Although particles smaller than 100 nm have been proposed as efficient carriers for organ-targeted delivery 62 , other studies suggest that appropriate surface functionalization can facilitate effective BBB translocation independent of size 63 . Our FA-NPs, composed of a ~110 nm silica core with a 220–260 nm and a low polydispersity index (PDI < 0.15), confirming uniformity and stable modification (Fig. 1C–E). The surface zeta potentials, ranging from −28.5 to −41.2 mV, are within the optimal range for stable nano-dispersions and BBB compatibility 64 (Fig. 1B). Anionic surface charges have been shown to better preserve BBB integrity and facilitate brain delivery compared to cationic particles 65,66 . Consistently, Zhang et al . and Chen et al . reported enhanced BBB penetration and brain accumulation of negatively charged mSiO 2 -NPs, with permeability influenced by both surface charge and particle size 65,67,68 . Moreover, PDA-coated mSiO 2 -NPs, but not uncoated silica particles, were recently shown to cross the BBB in vivo 47 . Our data align with these findings and support the conclusion that surface charge, controlled size, and functionalization collectively enable effective BBB permeability (Fig. 1B–E, Fig. 8). However, the relatively high hydrodynamic diameters (Fig. 1C–E) in sera might lead to enhanced accumulation of the particles in lung as previously reported and therefore require further investigation 60,69,70 . Mechanistically, our results revealed that FA-NP are internalized by microglia via FOLR-mediated, clathrin- and dynamin-dependent endocytosis (Fig. 2 D-J; Fig. 3 C-I). Uptake was significantly reduced by Dynasore and free FA, but unaffected by Cytochalasin D, ruling out phagocytosis and macropinocytosis. The significant uptake reduction by free FA in microglia, but not in astrocytes or neurons, highlights cell-specific folate receptor engagement. The nanoparticle size (~ 140 nm) further supports this pathway, as particles < 200 nm preferentially undergo clathrin-mediated uptake 71 , while caveolae-mediated internalization typically dominates above 500 nm and is less relevant in brain tissue due to low caveolin-1 expression in microglia 72 . While FOLR1 has been described preferentially upregulated by choroid plexus ependymal cells, oligodendrocyte precursor cells and midbrain dopamine progenitor cells 73–76 , FOLR2 has been found upregulated in embryonic meningeal border-associated macrophages and microglia 77 . More recent studies using spatial transcriptomics analysis reported FOLR2 upregulation in human adult microglia, suggesting a potential shift in receptor expression with maturation or disease context 78–80 . Supporting these transcriptomic observations, our own IHC and Western blot analyses confirmed the presence of both FOLR1 and FOLR2 proteins in human microglial cells, with a marked increase in abundance following LPS stimulation (Fig. 2). This LPS-induced upregulation may further enhance the selective uptake of FA-functionalized nanoparticles under inflammatory conditions. The preferential uptake we observed may therefore reflect a previously underappreciated translational window for targeting microglia in the adult brain via folate-linked delivery (Fig. 2A-C, Fig. 3B). Rapid accumulation in microglial populations within 1 minute of exposure, particularly in acute neuroinflammatory hotspots, highlights the spatial and temporal precision of this delivery platform (Fig. 3B, Fig. 4B-D). Once internalized, FA-NPs accumulated in endolysosomal compartments with extended retention over 96 h, accompanied by gradual exocytosis (Fig. 2A-C, Fig. 4E&F, Fig. 5A). The PDA coating conferred pH-responsiveness, as shown by time- and pH-dependent degradation of the polymer shell (Fig. 5A-C) and release of the encapsulated nuclear dye DAPI (Fig. 5D). TEM and ICC data (Fig. 5) suggest that full shell degradation is not required for cargo delivery, underscoring the utility of PDA for temporally controlled, intracellular drug release. Such behavior is critical in inflammatory environments characterized by acidic pH 81 . To achieve a sustained and effective therapeutic effect, FA-NPs must enable prolonged intracellular retention and controlled release of their payload. This typically requires successful escape from endolysosomal compartments, allowing cytoplasmic access to the cargo 82–84 . Our data suggest that PDA-coated NPs remain partially intact intracellularly over extended periods, as evidenced by the gradual degradation of the PDA shell observed up to 96 h post-incubation (Fig. 5A-C). This slow degradation supports a model of progressive intracellular release. Although we did not directly investigate the mechanism of endosomal escape, previous studies have proposed several possibilities for nanoparticle-mediated cytoplasmic delivery, including endosomal fusion, osmotic lysis via the proton sponge effect, vesicular rupture, and pH-triggered disassembly 85,86 . We also observed partially degraded FA-NPs in cell culture supernatants, indicating that a fraction of the nanoparticles is eventually exocytosed. Whether these extracellular particles can be taken up again by neighboring cells in multicellular systems or tissues remains unclear. Likewise, the fate of these particles in vivo , particularly their renal and hepatobiliary clearance dynamics, represents an important open question. Understanding the mechanisms governing systemic elimination and potential recirculation will be essential for advancing this platform toward safe and clinically viable applications. Surface functionalization is known to improve the biocompatibility of mSiO 2 -NPs by reducing nonspecific protein adsorption, cytotoxicity and hemolytic activity 60,87–89 . Additionally, NP-surface modifications with PEG or PDA have been shown to reduce uptake by the liver, spleen, and lungs, thereby prolonging systemic circulation and enhancing in vivo distribution profiles 65,90–92 . Our PDA- and FA-coated NPs exhibited reduced toxicity, ROS production, and cytokine expression compared to bare mSiO 2 -NPs (Fig. 6A–E). Encapsulation of CytoD within FA-NPs preserved its immunosuppressive effects, reducing microglial migration, phagocytosis, and cytokine production, while minimizing toxicity associated with free CytoD (Fig. 6F–I). In both hypoxia-reoxygenation and focal thermocoagulation injury models in adult murine brain slices, FA-NP[CytoD] treatment significantly suppressed IL-1β, IL-6, and TNF expression up to 96 h (Fig. 7). These results underscore the therapeutic potential of our system in modulating acute neuroinflammation. Notably, FA-NPs alone had no detectable immunosuppressive effects, confirming that therapeutic activity was cargo-dependent. This is consistent with previous findings that nanoparticle encapsulation enhances CytoD efficacy and safety in vivo 93 . Overall, these findings indicate that encapsulated CytoD retains its biological activity while targeted delivery circumvents systemic toxicity 27 , a key challenge that has historically limited its translational potential. Importantly, while many nanoparticle formulations face translational hurdles due to poor CNS delivery or off-target toxicity, our FA-NP[CytoD] system addresses both. Its modular design allows for tuning of surface chemistry, ligand density, and drug payload, while dual radiolabeling with 68 Ga and 177 Lu supports both diagnostic imaging and therapeutic tracking, key requirements for precision medicine. However, several limitations remain. Organotypic brain slices do not fully replicate the vascular and immune dynamics of the intact CNS, and LPS-induced microglial activation captures only part of the complex neuroinflammatory response after acute brain injury. Additionally, CytoD’s actin-modulating effects may differentially affect cell types and require longer-term evaluation in vivo . Future work should validate this platform in animal models of stroke or traumatic injury, assess long-term safety and efficacy, and explore disease-specific payloads tailored to microglial phenotypes. Additionally, optimization of dosing regimens, surface ligand density, and clearance kinetics will be essential for clinical translation. In summary, we present a modularly designed FA-NP[CytoD] platform that achieves BBB penetration, microglial targeting, controlled drug release, and real-time imaging capability. By integrating immunomodulation with therapeutic precision, FA-NP[CytoD] offers a promising strategy for the treatment of acute brain injury and potentially broader applications in neuroinflammatory and neurodegenerative diseases. These findings warrant further in vivo validation to advance this approach toward clinical translation. 4. Experimental Section Nanoparticle Synthesis and Functionalization Template-assisted Sol-Gel Synthesis of mSiO 2 -NPs : For the synthesis, an experimental description 88 was followed with minor modifications as previously described 51 . Initially, 0.365 g [1.0 mmol] CTAB was dissolved in 100 mL of water under magnetically stirring at an elevated temperature of 80 °C followed by the addition of 0.350 g NH 4 F. Afterwards, 1.80 mL TEOS was added dropwise to the clear solution with a speed of 1 mL/min and the mixture was stirred for another 2.5 h at 80 °C. After cooling down, the bluish-shimmering, white dispersion was centrifuged to collect mSiO 2 NPs. Subsequently, NPs were re-dispersed in acetone/diluted HCl and sonicated for 30 min to ensure the removal of any residuals of CTAB. Finally, additional washing steps of NPs with EtOH to remove any excess of released surfactant and silica were performed. Drying 24 h at ambient temperature, prepared the mSiO 2 -NPs for further characterization studies. Synthesis of Polydopamine-coated mSiO 2 -NPs : mSiO 2 -NPs were coated with polydopamine (PDA) by polymerizing dopamine under alkaline conditions. Therefore, a protocol 94 was followed with minor modifications as previously described 51 . Firstly, 0.040 g of mSiO 2 -NPs were dispersed in 60 mL of TRIS buffer (pH 8.5) [10 mM], and then 0.110 g dopamine hydrochloride was dissolved in the dispersion. The pH was adjusted again to pH 8.5 and the reaction was magnetically stirred for 24 h in the dark being exposed to oxygen. After adding dopamine hydrochloride, the reaction mixture immediately started to change its color turning from pale-yellow over yellowish orange to dark brown. The dark brown core-shell NPs were collected via centrifugation and washed several times with EtOH and water to remove any excess of polydopamine. Drying 24 h at ambient temperature, prepared the PDA-NPs for further characterization studies. Conjugation of Folic Acid on PDA-coated NPs : Folic acid was conjugated with its carboxy functions to residual NH 2 -groups of PDA-coated NPs via carbodiimide coupling reaction 95 with minor modifications as previously described 51 . For this approach, 0.130 g FA was dissolved in 100 mL acetone, 0.060 g of DCC was added and the reaction mixture was allowed to stir magnetically continuously at ambient temperature in the dark for 5 h. Afterwards, 0.035 g of NHS was added and the reaction mixture stirred at ambient temperature for further 24 h (in the dark). After pre-activation of the carboxy functions of FA with DCC and NHS, 0.105 g of PDA-NPs were re-dispersed in the reaction mixture carefully and the dispersion was stirred at ambient temperature in the absence of light for 48 h. Finally, the FA-NPs were separated from the supernatant via centrifugation and washed additional times with EtOH until any excess of FA was removed and the supernatant was colorless. Drying at ambient temperature for 24 h made the FA-NPs eligible for further characterizations. Chelator/Dye Attachment : To covalently attach DOTA as a chelator for radiometal labeling, as previously described 51 10 mg of either FA-NP or PDA-NP were re-dispered in 2 mL of a (1:1) Na 2 CO 3 -buffer/ultrapure water solution (Merck, Darmstadt, Germany). 2.9 mg of p-SCN-Bn-DOTA (Macrocyclics, Plano, USA) was dissolved in 2 mL Na 2 CO 3 -buffer (pH 9) and added dropwise to NP dispersion. The reaction mixture was heated up to 37 °C and stirred overnight. After cooling down, the NPs were collected via centrifugation (4000 rpm) and washed one time with Na 2 CO 3 -buffer and one time with ultrapure water before drying at ambient temperature. To covalently bind FITC, 400 µl of an aqueous FITC solution [1 mg/mL] (Merck, Darmstadt, Germany) was added to 1 mL of an aqueous FA-NP or PDA-NP dispersion [1.5 mg/mL] in metal free water. The mixture was stirred 6-8 hours in the dark. To remove unbound FITC the mixture was washed several times with ethanol/water (1:1) and centrifuged at around 4500 x g for 10 min and the supernatant was removed. The washing procedure was repeated up to six times until supernatant is colorless. Drug Loading : Drug loading of Cytochalasin D (CytoD) or 4′,6-Diamidino-2-phenylindol (DAPI) into mesoporous NPs was achieved via a diffusion effect. To 250 µl of an aqueous NP dispersion [1.5 mg/mL], 0.5 µl [5 mg/mL EtOH] CytoD (Merck, Darmstadt, Germany) or 5 µl [0.02 mg/mL H 2 O] DAPI (Roth, Karlsruhe, Germany) was added, and the mixture was magnetically stirred for 24 h at RT. For Purification the mixture was centrifuged (4000 rpm for 5 min) and washed three times with ultrapure water (Merck, Darmstadt, Germany) while the supernatant was collected. Drug Release : To determine the pH-dependent PDA-shell degradation, NPs were incubated extracellular in media for 48 h at pH values of 7.4 or 5.0. Furthermore, to evaluate the intracellular PDA degradation FA-NPs were incubated for 1 h in microglial HMC-3 cells and were assessed after retention times of 48 h or 96 h. TEM images were taken, and ImageJ was used to analyze the PDA mean thickness r (four different points for each NP) and the PDA mean area in all four conditions. Nanoparticle Characterization Zeta Potential : The zeta potential values (ζ-potential) of the bare and surface engineered NPs were measured on a Zetasizer Nano ZS (Malvern Panalytical, Malvern, UK) in aqueous solution in triplicate. Fourier-Transform Infrared spectroscopy : To evaluate the successful NP synthesis, modification and functionalization, Fourier-Transform Infrared (FT-IR) spectra for the detection of functional groups on the synthesized and functionalized NPs were recorded as previously shown 51 on a Spectrum 400 (PerkinElmer, Waltham, USA) with Universal ATR sampling accessory in the range of 4000 to 400 cm -1 . Transmission Electron Microscopy : For morphology, size and shape analysis the NPs were were fixed in 3% glutaraldehyde in 0.1 M Soerensen’s phosphate buffer (Merck, Darmstadt, Germany) and embedded in 2.5% low-melting agarose (Sigma, Steinheim, Germany). After post-fixing with 1% OsO4 (Roth, Karlsruhe, Germany) in 25 mM sucrose buffer (Merck, Darmstadt, Germany) the samples were dehydrated by ascending ethanol series (30, 50, 70, 90 and 3 times 100%) for 10 min each. Dehydrated samples were incubated in propylene oxide for 30 min., then in a mixture of Epon resin (Serva, Heidelberg, Germany) and propylene oxide (Science Services, Munich, Germany) (1:1) for 1h and finally in pure Epon for 1h. Samples were embedded in fresh Epon and polymerized at 90°C for 2h. Ultrathin sections (~90 nm) were stained with 0.5% uranyl acetate and 1% lead citrate (both EMS, Munich, Germany) to enhance contrast. Samples were examined using a transmission electron microscope (Zeiss Leo906, Oberkochen, Germany) operating at an acceleration voltage of 60 kV. Nanoparticle Tracking Analysis : Size distribution and polydispersity of the NPs were determined using nanoparticle tracking analysis (NTA) measurements on a LM10SH (NanoSight, Malvern, United Kingdom), equipped with a sample chamber with a 532 nm laser. The captured videos were analyzed by the NTA 2.0 image analysis software (NanoSight, Malvern, UK). The mode and mean size and SD values were obtained by the NTA software. Radiochemistry Radiolabeling : Radiolabeling was carried out as previously described 51 . To 1.5 mg of p-SCN-Bn-DOTA-modified FA-NP or PDA-NP 1 mL metal free ultrapure water (Merck, Darmstadt, Germany) was added and the mixture was sonicated for 5 min at 30 °C. 40-250 µl [1.5 mg/mL] of that NP dispersion were transferred to a 2 mL glass reaction vial, using an aqueous ammonium acetate solution [3 M] as buffer . In case of 68 Ga-labeling, up to 110 µl [ 68 Ga]GaCl 3 (up to 500 MBq) were added. The radionuclide was supplied by a 68 Ge/ 68 Ga-generator (iThemba, Cape Town, South Africa) using 0.6 M HCl as eluent. In case of 177 Lu-labeling, up to 1 GBq 177 LuCl 3 (volume 98 %) and radiochemical purities (> 98 %) of 68 Ga- and 177 Lu-labeled NPs were determined by radio-TLC (Elysia-Raytest, Straubenhardt, Germany) with citrate buffer (pH 5.0) as mobile and iTLC-SG strips (Agilent, Santa Clara, USA) as stationary phase. In vitro Stability : Stability of 68 Ga and 177 Lu-labeled NPs was determined as previously described 51 , by incubating the radiolabeled compound in PBS (pH 7.4) or human sera (1:9) at 37 °C. Aliquots thereof were analyzed at specified time points (for 68 Ga at: 15, 45, 60, 90, 120 and 180 min; for 177 Lu at: 0, 1, 2, 4, 24, 48, 72, 96 and 120 h) by radio-TLC regarding the present fraction of labeled NPs. Human sera were obtained from three healthy adult female and three healthy adult male donors. Partition Coefficient : As previously described 51 , to a 1:1 water/hexane mixture, 2 MBq of 177 Lu-labeled NPs were added. The mixture was vigorously vortexed for at least 10 min. 0.1 mL of each layer was pipetted into a test tube and the radioactivity was measured using a Wizard 2 gamma counter (PerkinElmer, Waltham, USA). The measurement was carried out in triplicate. The partition coefficient (P) was calculated as a ratio of counts in the hexane phase to counts in the PBS phase (background corrected) and logarithmized to yield the logP. Cells Human HMC-3 microglial cell line (ATCC: CRL-3304), were kindly provided by Prof. Cordian Beyer (Institute of Neuroanatomy, Medical Faculty, RWTH Aachen University). Human glioblastoma astrocytoma cell line U-373 MG (Uppsala) (ECACC 08061901) were kindly provided by Dr. Brunkhorst (Department of Neurology, Medical Faculty, RWTH Aachen University). Human neuroblastoma cell line SH-SY5Y was politely provided by Dr. Aaron Voigt (Department of Neurology, Medical Faculty, RWTH Aachen University). HMC-3, U-373 and SY5Y cells were cultivated in Dulbecco’s modified Eagle’s medium (DMEM, Pan Biotech, Aidenbach, Germany) supplemented with 10 % fetal bovine serum (FBS, Pan Biotech, Aidenbach, Germany) and 0.5 % penicillin/streptomycin (P06-07100 (P/S, Pan Biotech, Aidenbach, Germany)). All cells were maintained in a humidified incubator at 37 °C and 5 % CO 2 . HMC-3 cells were splitted every 2 days at approximately 75 % confluency by trypsinization (Trypsin/EDTA, Pan Biotech, Aidenbach, Germany). Regular PCR controls against gene markers of cell types used in our laboratory ensured the purity of the cell lines. One day prior to experiments the cells were detached by trypsinization with subsequent centrifugation (400 g, 5 min, room temperature (RT)). The cell pellet was resuspended in DMEM, 5 % FBS, 0.5 % P/S (all Pan Biotech, Aidenbach, Germany) and seeded in 6-well dishes for experiments and on coverslips for ICC. Cell Uptake Transmission Electron Microscopy : Microglia (HMC-3), Astrocyte (U373) and Neuron (SY5Y) cell lines were cultured in 6-well plates with a seeding density of 750x10 3 cells/well. To assess the nanoparticle uptake, individual well was treated with a cold FA-NP formulation diluted in 3 mL culture media and incubated for 1 hour at 37 °C. After washing with PBS, the cells were fixed in 3% glutaraldehyde in 0.1 M Soerensen’s phosphate buffer (Merck, Darmstadt, Germany), scratched off the plate, pelleted via centrifugation and embedded in 2.5% low-melting agarose (Sigma, Steinheim, Germany). Samples were further processed for transmission electron microscopy as already described. Gamma Counter : To assess the quantitative nanoparticle uptake, as previously described 51 individual well was treated with different 177 Lu-labeled nanoparticle formulations of either functionalized with folic acid or control in a < 0.01 Bq/cell concentration diluted in 3 mL culture media and incubated for 1 hour at 37 °C. After washing with PBS, the cells were trypsinized, collected in a γ-Counter test tube, and their radioactivity was measured using a γ-Counter (Wizard 2 , PerkinElmer, Waltham, USA) with a protocol for 177 Lu calibration. Efficiency corrected cell total activities were then expressed as counts per minute. Each experiment was performed three times with three technical replicates respectively. To determine the percentage of NP incorporation into the cells, the exact total initial activity of NP was calculated by counting 1 mL of a 1:10 dilution after the same incubation time. Background radiation was determined with 1 mL of culture medium. The percentage of NP incorporation was calculated using the following formula: \begin{equation} \text{NP}_{I}\left[\%\right]=\frac{A_{\text{cells}}-A_{\text{Background}}}{A_{0}}\times 100\nonumber \\ \end{equation} , where NP I is the incorporation percentage and A cells , A Background , A 0 are the disintegrations per minute, registered for the cells, background radiation, and the dilution-corrected initially activity, respectively. To determine the relative NP uptake of each condition, the γ-Counter measured uptake was calculated as percentage from the initial 177 Lu-labeled nanoparticle-culture media solution. For uptake studies with inhibitor treatment, cells were pretreated for 30 minutes with free FA [1 mM], CytoD [10 µM], Dynasore [10 µM], Rottlerin [6 µM], or a combination of free FA [1 mM] + Dynasore [10 µM] or free FA [1 mM] + Rottlerin [6 µM] prior to nanoparticle addition for an incubation period of 1 h at 37 °C. Western Blot Western blot (WB) analysis was performed as previously described 96,97 . After incubating microglial HMC-3 cells with nanoparticles, cells were lyzed ice-cold radioimmunoprecipitation assay buffer (RIPA) supplemented with phosphatase and proteinase inhibitors (cOmplete, #1187358001, Roche, Basel, Switzerland). 20 µg of protein was loaded onto the SDS-PAGE gel and separated. After the transfer onto the PVDF membrane (Roche, Basel, Switzerland) and blocking with 5 % skim milk (#T145.3, Roth, Karlsruhe, Germany) in tris-buffered saline containing 0.05 % Tween20 (#8076.4, Roth, Karlsruhe, Germany), we incubated the membrane with the primary antibody in corresponding concentrations (Table 1). Following the incubation with the secondary horseradish-conjugated antibody, we visualized the results using ECL™ Plus Kit (Thermo Fisher Scientific, Waltham, MA, USA). β-Actin served as a loading control. Densitometric analysis was performed using ImageJ (NIH, Bethesda, MD, United States). Table 1. List of antibodies used for Western Blot. FolR1 Cloud-Clone Corp., TX, USA PAF460Ra01 Rat 1:500 FolR2 Abcam, Cambridge, United Kingdom Ab103998 Rabbit 1:500 Rabbit IgG GE, Healthcare, Chicago, IL, USA NA934 Goat 1:5000 Mouse IgG GE Healthcare, Chicago, IL, USA NXA931 Goat 1:5000 β-Actin Abcam, Cambridge, United Kingdom Ab3280 Mouse 1:1000 Immunocytochemistry (ICC) ICC analysis was performed according to established protocol 96,97 . Cells were seeded to Poly-L-lysine covered cover slips. After stimulation, cover slips were fixed with 4 % formaldehyde in phosphate buffered saline (PBS) and permeabilized with 0.2 % Triton X-100. Subsequently the slides were blocked with Immunofluorescence Blocking Buffer (1 % BSA, 2 % FBS in PBS) and incubated with the primary antibody overnight at 4 °C. The secondary antibody was incubated for 1 h at room temperature. Negative controls were incubated with only PBS buffer overnight. For the time-dependent intracellular retention time and drug release experiment of FITC-FA-NPs in microglia, LysoTracker TM Red DND-99 (Thermo Fisher Scientific, Waltham, MA, USA) was added 1 h before the end of nanoparticle incubation time. Cell nuclei were stained with 4′,6-Diamidino-2-phenylindol (DAPI) (Roth, Karlsruhe, Germany). Stainings were evaluated using a Leica fluorescence microscope (Leica, Wetzlar, Germany). List of antibodies used for immunocytochemistry is given in Table 2. Table 2. List of antibodies used for immunocytochemistry and immunohistochemistry. IBA1 Wako Chemicals, Neuss, Germany 019-19741 rabbit 1:1000 GFAP Millipore, Burlington, MA, USA MAB3402 mouse 1:500 NeuN Millipore, Burlington, MA, USA MAB377 mouse 1:1000 goat anti-mouse IgG Alexa Fluor 594 Invitrogen, Karlsruhe, Germany A11032 goat 1:500 goat anti-rabbit IgG Alexa Fluor 594 Invitrogen, Karlsruhe, Germany A11012 goat 1:500 Fluorescence Activated Cell Sorting HMC-3 cells were incubated with Rottlerin [6 µM], Dynasore [10 µM], CytoD [10 µM], CytoD-loaded FA-NPs [6.25 µg/mL] and H 2 O 2 (0.1 %) for 1 h at 37 °C. After 1 h of incubation time with the mentioned treatments, medium was changed, and cells were maintained for 24 h. After 24 h, cells were harvested by taking the supernatant and incubating the remaining cells for 5 min with trypsin (at 37 °C). HMC-3 cells were centrifuged and stained with Annexin V (Alexa Fluor 647) and PI and analyzed with FACSCanto™ II (BD Bioscience, Erembodegem, Belgium). Cell Viability Cell viability was measured as previously described 96 , using a lactate-dehydrogenase (LDH) assay according to the manufacturer’s protocol (LDH, CytoTox 96 ® Non-Radioactive Cytotoxicity Assay, Promega, Walldorf, Germany). The release of stable cytosolic LDH into the extracellular space occurs during late apoptosis or necrosis of cells. Therefore, the amount of released LDH was measured in the supernatant at 490-520 nm using a microplate reader (Tecan, Männedorf, Switzerland). Cells treated with lysis solution served as an internal positive control, as previously shown 96,98 . The percentage of released LDH was determined by referring to the positive control, which was referred to as 100 % of possible LDH release. The findings were validated utilizing Trypan Blue staining (#T8154, Sigma-Aldrich, St. Louis, USA). Here, viable cells were counted by an automated cell counter (Roche Innovatis Cedex XS, Basel, Switzerland). Metabolic Activity Cell metabolism was measured as previously described 96 , by CellTiter-Blue assay according to the manufacturer’s protocol (CellTiter-Blue assay, Promega, Walldorf, Germany). The reduction of blue resazurin to pink, fluorescent resorufin requires an intact mitochondrial respiratory chain and can be observed only in metabolically active cells. The reduction of resazurin was observed in viable cells at 590 nm by a microplate reader (Tecan GmbH, Männedorf, Switzerland) as previously described 96,99 . ROS Assay The ROS assay was performed according to the manufacturer’s protocol and as previously described 96 . In brief, 5 µM CellROXTM (C10444, Thermo Fisher Scientific, Waltham, MA, United States) reagent was added to the medium. After 30 min of incubation at 37 °C the supernatant was removed, and the cells were washed three times with PBS. Fluorescence was measured at an excitation wavelength of 488 nm and emission wavelength of 525 nm as previously described 96 . Reverse Transcription Quantitative PCR Gene expression analysis was performed as previously described 96 with human microglial HMC-3 cells. After dissolving and homogenizing cells in PegGold (PeqLab #30-2010, Erlangen, Germany) total RNA was extracted using peqGold RNA TriFast as previously described 96,97 . Complementary DNA was synthesized using an iScript TM cDNA Synthesis Kit (Bio-Rad Laboratories, CA, United States) and random hexanucleotide primers (Invitrogen, Karlsruhe, Germany) using 1 µg of total RNA according to the manufacturer’s protocol. Triplicates of every sample were transferred by a pipetting robot (Corbett CAS-1200, Qiagen, Hilden, Germany) to Rotor-Gene strip reaction tubes (I1402-0400, StarLab, Hamburg, Germany) and RT-qPCR analysis was performed using the Rotor-Gene Q device (Qiagen, Hilden, Germany). RNase free H 2 O (Merck, 64293, Darmstadt, Germany) served as no template control (NTC) and primer efficiencies were calculated using the Pfaffl method 100 . The target genes and housekeeping gene (glyceraldehyde-3- phosphate dehydrogenase, GAPDH) were measured at cycle threshold (Ct values), and relative quantification was calculated by the 11Ct method using the qbase+ software (Biogazelle, Belgium). The following forward (fwd) and reverse (rev) primers were used (50 - > 3 0 ): GAPDH (fwd: CCT GCA CCA CCA ACT GCT TA, rev: GGCCATCCACAGTCTTCTCAG), IL-6 (fwd: ACTCACCTCTTCAGAACGAATTG, rev: CCATCTTTGGAAGGTTCAGGTTG), IL-1β (fwd: CTT CGA GGC ACA AGG CAC AA, rev: TTC ACT GGC GAG CTC AGG TA), TNF (fwd : CCTCTCTCTAATCAGCCCTCTG, rev: GAGGACCTGGGAGTAGATGAG), CCL2 (fwd: CAGCCAGATGCAATCAATGCC, rev: TGGAATCCTGAACCCACTTCT), CCL5 (fwd: CCAGCAGTCGTCTTTGTCAC, rev: CTCTGGGTTGGCACACACTT), BAX (fwd: CCCGAGAGGTCTTTTTCCGAG, rev: CCAGCCCATGATGGTTCTGAT), BCL2 (fwd: GGTGGGGTCATGTGTGTGG, rev: CGGTTCAGGTACTCAGTCATCC), CASP 3 (fwd : CATGGAAGCGAATCAATGGACT, rev : CTGTACCAGACCGAGATGTCA), CASP8 (fwd: TTTCTGCCTACAGGTTCCACT, rev: CCTCAATTCTGATCTGCTCACTT), CASP9 (fwd: CTTCGTTTCTGCGAACTAACAGG, rev: GCACCACTGGGGTAAGGTTT). Zone Exclusion/Cell Migraton Assay Zone exclusion assay was performed like previously described by Arik et al . 96 . For 48 h the migration of HMC-3 cells was observed. One day prior to the experiments 70 µl of 500,000 cells/mL were seeded into 2 well culture-inserts (80209, Ibidi GmbH, Germany). After 24 h the medium was changed, and cells were left to incubate for 1 h with different conditions. Treatment conditions included drug-unloaded FA-NP [6.25 µg/mL], FA-NP-encapsulated Cytochalasin D (#C2618, Sigma-Aldrich, Taufkirchen, Germany) [6.25 µg/mL] and molecular CytoD [5 µM] dissolved in DMEM with 5 % FBS, 0.5 % P/S. Medium with NaCl and only 5 % FBS and 0.5 % P/S served as vehicle control. Afterwards, the inlets were removed, thus creating a 500 µm sized gap between both cell monolayers and the medium was changed to 5 % RPMI with 0.5 % P/S and an additional Mitomycin C [10 µg/mL] (10107409001, Roche, Germany). Mitomycin C served as a proliferation inhibitor. Cells were monitored using live cell imaging (Carl Zeiss, Oberkochen, Germany). Measurements were repeated accordingly with different CytoD-loaded FA-NP concentrations [1.5, 3.12, 6.25, and 12.5 µg/mL] to evaluate the concentration-dependent migration. Furthermore, measurements were performed in comparing the migration using FA-NP[CytoD] + Dynasore [5 µM] versus drug-unloaded FA-NP + Dynasore [5 µM]. Phagocytosis Phagocytic activity of HMC-3 microglial cells was evaluated by measuring the uptake of pHrodo™ Red-conjugated E. coli bioparticles (pHrodo™ Red E. coli Bioparticles™, Thermo Fisher, Waltham, MA, USA), as previously described 96,98,101 . Cytochalasin D [5 µM] (#C2618, Sigma-Aldrich, Taufkirchen, Germany) was used as a negative control, whereas lipopolysaccharide [1 µg/mL] (from Salmonella minnesota, R596(Re) (TLRgrade™), ALX-581-008-L002, Enzo, Farmingdale, NY, USA) served as a positive control. Phagocytic activity was assessed at 24 h and 48 h following a 1-hour incubation with either NaCl (control), FA-NP [6.25 µg/mL], FA-NP[CytoD] [6.25 µg/mL], molecular CytoD [5 µM], or LPS [1 µg/mL] (positive control). For this, cells were incubated with pHrodo™ Red E. coli Bioparticles™ for 1 hour, followed by three washes with PBS medium. Quantification was performed by counting pHrodo™-positive cells per 1,000 cells in each experimental condition 96 . Organotypic Murine Brain Slice Cultures Coronal brain slices with a thickness of 350 µm were prepared using a vibrating-blade microtome (Leica VT1200 S) and maintained in ice-cold, carbonated artificial cerebrospinal fluid (aCSF) during processing. For cultivation, the slices were placed onto 30 mm Millicell-CM tissue culture inserts with 0.4 µm pore size (Millipore) positioned in six-well culture plates containing long-term aCSF. Cultures were then incubated at 37 °C with 5% CO 2 in a HERAcell cell culture incubator (Thermo Fisher). 350 µm formalin fixed organotypic adult murine brain slices were incubated in 10 % sucrose for several hours, followed by incubation in 30 % sucrose overnight. Subsequently, 12 µm coronal brain sections were prepared using a cryostat (Leica CM 3050S, Wetzlar, Germany) and directly mounted onto microscope slides. After washing in PBS, the slides were blocked with a blocking buffer (3 % normal goat serum in PBS) for 30 min. Then, they were incubated with the primary antibody, diluted in PBS (Table 2), overnight at 4 °C. The following day, the slides were washed in PBS and the secondary antibody (Table 2), diluted in PBS (1:500), was applied for 1 h at room temperature. Nuclear counterstaining was performed by incubating the slides in DAPI (4′,6-diamidino-2-phenylindole) for 1 min. After a final wash in PBS, the slides were mounted using Fluoromount-G. Brain Injury Models Hypoxia-Reoxygenation : To mimic ischemic stroke, we performed Oxygen-Glucose-Deprivation (OGD) as previously described 96 , using 350 µm thick coronal organotypic adulte murine brain slice cultures. Oxygen depletion was induced in our customized hypoxia chamber (Part#: C174, C21, BioSpherix, Parish, NY, United States) by flooding the chamber with 100 % inert nitrogen gas until the oxygen concentration reached < 1 %. To ensure stable OGD-conditions we monitored oxygen levels, temperature and pressure within the chamber. An additional oxygen sensor was used to monitor the oxygen level in the media (#200001735, PreSens, Regensburg, Germany). An orbital shaker was placed beneath the hypoxia chamber at 30 rpm ensuring better O 2 depletion in the media. The chamber itself was placed in a humidified incubator set to 37 °C. Normoxia controls were kept at 37 °C, 5 % CO 2 and atmospheric pressure. Focal Thermocoagulation : To mimic traumatic brain injury, we used a focal thermocoagulation injury model targeting the primary and secondary motor cortex, with involvement of both cortical and subcortical regions, including the corpus callosum using 350 µm thick coronal organotypic adulte murine brain slice cultures. Animal Experiments All animal experiments were approved by the appropriate German authorities (LANUV: Landesamt für Natur, Umwelt und Verbraucherschutz Nordrhein-Westfalen) in accordance with national regulations. Six NOD/SCID female mice were purchased from Janvier Laboratories (NOD.CB17-Prkdc scid/scid /Rj). The animals were housed in the Department of Nuclear Medicine of the University Hospital RWTH Aachen under a 12-h-light/12-h-dark cycle and were given ad libitum access to feed and water. The room temperature and relative humidity were kept between 20-25 °C and 45–65 %, respectively. In vivo Biodistribution Mice were imaged with a small animal SPECT/CT system (X-CUBE and γ-CUBE, Molecubes, Gent, Belgium) as previously described 51 . Under 1.5 to 2.5 % isoflurane anesthesia in oxygen at 0.8 L/min, the lateral tail vein was injected with 30 MBq of [ 177 Lu]Lu-DOTA-FA-NPs ( 177 Lu-FA-NPs) diluted in 0.9 % NaCl to a total volume of 125 μL and scanned 4 h or 24 h post-injection. CT acquisition settings: Default high resolution protocol, 440 µA, 50 kVp, and 32 ms exposure time; the duration of each CT scan was approx. 5 min. The SPECT scan was initiated at the end of the CT. SPECT acquisition settings used: 7-headed camera equipped with a general-purpose collimator with 28 pinholes (0.75mm apertures and a radius of rotation of 20 mm) and a total duration of 30 min. During the scans, the isoflurane concentration was adapted to achieve a respiratory rate between 75 - 50 breaths per min. CT images were reconstructed as previously described 51 , using an iterative reconstruction algorithm (ISRA) reconstruction process to an isometric voxel size of 0.2 mm in a 200 × 200 × 425 matrix. Using a vendor software, CT values were converted into Hounsfield units (HU) using the formula: HU = 1000 × ((µ t − µ w ) / (µ w - μ a ), where µ w is the linear attenuation coefficient of water, µ a is the linear attenuation coefficient of air water and µ t is the linear attenuation coefficient of the tissue. SPECT data were reconstructed using a 3-dimensional maximum likelihood estimation method (MLEM-3D with 50 iterations) into a 141 × 141 × 425 image matrix (resulting in final isometric voxel size of 0.25 mm) with an energy window of ± 10 % from a peak of 113 keV and of 210 keV. SPECT and CT images were automatically aligned after reconstruction. PMOD software package version 3.13 (PMOD Technologies LLC, Zürich, Switzerland) was used to view, render, and analyze the SPECT/CT data. For the 3D rendering of the brain soft tissue, the inner side of the skull and vertebral canal was auto-segmented using a CT-based thresholding at 700 HU. Afterwards the outside of the auto-contour was masked, while the inside was used for 3D rendering using the P3D tool. The IN RANGE segmentation method was applied with -100 HU and 700 HU as limits, resulting in a single volume rendered object of the brain soft tissue. The co-registered SPECT acquisition was used as texture, with 0 and 800 prop. to cnts/sec as limits, using a Hot color scale. For the maximal intensity projection images (MIP), the PFUS tool was used with the following settings: for CT, the limits were set to 1 % and 100 % of the minimal and maximal HU value, while for SPECT the limits were set at 0 and 3100 prop. to cnts/sec. Brains (24 h or 120 h p.i.) were perfused with PBS to remove residual blood, fixed, harvested and cut into 8 (2 mm thick) coronal brain sections. They were collected in γ-Counter test tubes, and their radioactivity was measured using a γ-Counter (Wizard 2 , PerkinElmer, Waltham, USA) with a protocol for 177 Lu calibration. Radioactivity was then expressed as counts per minute (CPM). Thereafter, the coronal brain sections were placed on imaging plates (Fuji Film BAS-IP SR 2025, Raytest, Germany) for 24 h at room temperature and subsequently scanned using the Typhoon FLA 7000 (GE Healthcare, IL, USA) to visualize the 177 Lu-FA-NPs distribution by ARG prior to IHC analysis of FITC-FA-NPs in the same brain sections (Table 2). Statistics We utilized GraphPad Prism (version 8.4.3, San Diego, USA) for data analysis and visualization. Residuals were analyzed for normal distribution using the Shapiro-Wilk and D ’ Agostino-Pearson omnibus normality test. Variance homogeneity was tested using the Bartlett test or the Spearman ’s rank correlation test for heteroscedasticity. We utilized a ROUT test to identify outliers. P < 0.05 was considered statistically significant. Detailed description of biological and technical numbers are given in the corresponding figure legends including the statistical tests employed. Author Contributions SEMS.: Writing, original draft, visualization, methodology, investigation, formal analysis, data curation. RN and PA: Investigation, data curation, review and editing. JP, LG, SI, EA, AF, EMB: Investigation, validation, review and editing. BN: Methodology, review and editing. SM, JBS, FMM: Resources, review and editing. PH: Review and editing, investigation, supervision, resources, methodology, funding acquisition, formal analysis, conceptualization, project administration. Data Availability Statement All data supporting the findings of this study are available within the article and its supplementary information files. Raw data and additional information related to this paper may be requested from the corresponding author upon reasonable request. Acknowledgements and Funding We thank Sabine Hamm, Fiona Teubner (Department of Neurology, Medical Faculty, RWTH Aachen University) for their excellent technical assistance. We thank Annika Szymura (Institute of Inorganic and Materials Chemistry, University of Cologne) for her valuable assistance in nanoparticle synthesis. We thank Patrick Robert Buchta from the Audiovisual Media Center (AVMZ), Medical Faculty, RWTH Aachen University, for his support with the graphical visualization of the nanoparticles. We thank the workgroup of Prof. Jahnen-Dechent (Helmholtz Institute for Biomedical Engineering, Biointerface Laboratory, RWTH Aachen University) for their technical support in the NTA analysis. This work was funded by an internal grant (START grant 111/17, PH) and the German Research Foundation (DFG), HA 9566/1-1 (PH) and supported by the “Clinician Scientist program” of the Faculty of Medicine, RWTH Aachen University (PH). 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Keywords neurotherapeutics precision medicine targeted drug delivery Authors Affiliations Sabri Elyes Mohamed Sahnoun Universitatsklinikum Aachen View all articles by this author Rozina Noristani Stanford University School of Medicine View all articles by this author Paolo Alimonti University of Oxford Department of Oncology View all articles by this author Jonas Pes Universitatsklinikum Aachen View all articles by this author Lara Gubeljak Universitatsklinikum Aachen View all articles by this author Shaista Ilyas Universitat zu Koln Department fur Chemie View all articles by this author Eren Arik Universitatsklinikum Aachen View all articles by this author Alexandru Florea Universitatsklinikum Aachen View all articles by this author Eva Miriam Buhl Universitatsklinikum Aachen View all articles by this author Bernd Neumaier Forschungszentrum Julich GmbH Projekttrager Julich View all articles by this author Sanjay Mathur Universitat zu Koln Department fur Chemie View all articles by this author Jörg B. Schulz Universitatsklinikum Aachen View all articles by this author Felix Manuel Mottaghy Universitatsklinikum Aachen View all articles by this author Pardes Habib 0000-0002-5771-216X [email protected] Stanford University School of Medicine View all articles by this author Metrics & Citations Metrics Article Usage 343 views 163 downloads .FvxKWukQNSOunydq8rnd { width: 100px; } Citations Download citation Sabri Elyes Mohamed Sahnoun, Rozina Noristani, Paolo Alimonti, et al. Targeted nanocarriers for microglia-selective cytochalasin D delivery to modulate neuroinflammation after acute brain injuries. Authorea . 18 November 2025. DOI: https://doi.org/10.22541/au.176349281.18872463/v1 If you have the appropriate software installed, you can download article citation data to the citation manager of your choice. Simply select your manager software from the list below and click Download. For more information or tips please see 'Downloading to a citation manager' in the Help menu . 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