This article mainly describes a fermentation and purification method for expressing recombinant collagen protein in Escherichia coli. The method comprises constructing engineered bacteria expressing human type III collagen and adopting a strategy of feeding in batches for high-density fermentation. The rapid proliferation of bacterial cells is promoted at 37°C, and then the culture is inoculated into a fermentation tank with different carbon sources for growth. When glycerol is used as the main carbon source, the yield of recombinant collagen protein can reach 0.25 to 0.40 g/L. The method allows exploration of the differences in recombinant collagen production with different carbon sources in order to identify the most suitable fermentation medium component. The human type III collagen produced by the method has the typical structure of collagen, with high cell adhesion and the stability of tissue structure. Therefore, it can be used as the raw material for various collagen products, especially facial fillers, dressings, freeze-dried fibers, and gels. © 2025 Wiley Periodicals LLC. Basic Protocol: Fermentation and purification of recombinant human type III collagen expressed in Escherichia coli.

DNA replication is often challenged by endogenous and exogenous sources of DNA damage, which can stall replication forks and result in single-stranded DNA (ssDNA) gaps, double-strand breaks (DSBs), and genomic instability. Detecting DNA damage specifically in newly synthesized DNA strands is essential for understanding how eukaryotic cells respond to replication stress and continue the cell cycle progression through DNA repair or DNA damage tolerance (DDT) mechanisms. Here, we present an optimized and accessible protocol for the alkaline BrdU comet assay-a single-cell technique that combines bromodeoxyuridine (BrdU; a thymidine analog) pulse-labeling of newly synthesized DNA with the alkaline comet assay (single-cell gel electrophoresis) followed by fluorescence immunodetection. This method enables the specific detection and measurement of DNA strand breaks occurring in newly replicated DNA during and immediately after the S phase, even without cell synchronization, allowing researchers to differentiate replication-associated DNA damage from overall genomic damage. We provide detailed instructions for performing the assay using human cells (RPE-1 h-TERT TP53 KO) in vitro after exposure to DNA replication-stress-inducing agents, such as hydroxyurea (HU) and ultraviolet-C (UV-C) radiation. We also demonstrate its application in translesion-synthesis-deficient (Pol eta-deficient) human fibroblasts in vitro. Importantly, this protocol supports time-course chase experiments (e.g., 0, 1, 2, and 4 hr post-treatment) to monitor the kinetics of DNA damage in nascent DNA strands. This BrdU-based protocol offers high specificity, single-cell resolution, and cost-effectiveness, making it particularly valuable for laboratories studying replication stress, post-replication DNA repair proficiency, DDT, and/or genotoxic responses in unsynchronized human cells in vitro. This protocol also adheres to the Minimum Information for Reporting Comet Assay (MIRCA) guidelines and is aligned with the objectives of the International Comet Assay Working Group (ICAW), ensuring high reproducibility and standardization. © 2025 The Author(s). Current Protocols published by Wiley Periodicals LLC. Basic Protocol: Alkaline BrdU comet assay to assess replication-associated DNA damage in unsynchronized human cells (RPE-1 h-TERT TP53 KO) in vitro Alternate Protocol 1: Alkaline BrdU comet assay to monitor replication-associated DNA damage dynamics in unsynchronized human cells (RPE-1 h-TERT TP53 KO) after HU and UV-C exposure and 0- to 2- hr chase Alternate Protocol 2: Alkaline BrdU comet assay to compare replication-associated DNA damage dynamics in translesion synthesis polymerase η-deficient and complemented (XP-V comp) unsynchronized fibroblasts after UV-C exposure and 0- to 4-hr chase.

Biomaterial‐based bioinks are increasingly utilized in bioprinting to engineer three‐dimensional (3D) constructs with living cells for tissue engineering and disease modeling. Among various bioinks explored, alginate‐based formulations stand out due to their good biocompatibility, mild gelation conditions, tunable mechanical properties, and ease of crosslinking via divalent cations such as calcium. Despite their widespread use, standardized protocols for preparing alginate‐based bioinks and characterizing bioprinted constructs have not been well documented. Our laboratory has developed and validated reproducible methods for preparing a variety of alginate‐based bioinks and printing cell‐laden constructs tailored for diverse applications. In this article, we present detailed step‐by‐step protocols covering bioink preparation and rheological characterization, extrusion‐based bioprinting of cell‐laden constructs, post‐printing culture and co‐culture techniques, printability assessment, and live/dead and immunofluorescence assays. These protocols serve as a standardized framework for the fabrication and characterization of 3D bioprinted alginate‐based cell‐laden constructs, thereby facilitating translational research in tissue engineering, disease modeling, and preclinical therapeutic development. © 2025 The Author(s). Current Protocols published by Wiley Periodicals LLC.Basic Protocol 1: Bioink preparationBasic Protocol 2: Bioink characterization using rheologyBasic Protocol 3: Scaffold design and bioprintingSupport Protocol: 3D‐printing parameter determinationBasic Protocol 4: Printability and cell viability analyses, and immunofluorescence assay

The aim of this study was to develop an improvised protocol for ex vivo expansion of caprine satellite cells (SCs) using growth factors and their cryopreservation. The SCs were isolated from muscle tissue using collagenase type-I for enzymatic digestion. In vitro culture of SCs was done using Dulbecco's modified Eagle medium (DMEM) supplemented with 20% fetal bovine serum (FBS) and 15% horse serum. Immunofluorescence and PCR confirmed positive expression of PAX7 and MyoD, and negative expression of MyHC. Supplementation with 10 ng/ml basic fibroblast growth factor (bFGF) + 100 ng/ml insulin-like growth factor-1 (IGF-1) in the basal culture medium (DMEM + 2% FBS) showed similar results as control (routine culture medium). Subsequently, SCs were successfully cryopreserved using 10% dimethyl sulfoxide and 40% FBS with post-thaw viability of 74.72%. We have successfully established a protocol for the isolation, culture, and cryopreservation of goat SCs. Caprine SCs can be successfully expanded in vitro using growth factors (10 ng/ml bFGF and 100 ng/ml IGF-1) with minimum supplemental FBS in basal culture medium. Additionally, investigations could assess the viability and integrity of cryopreserved goat SCs post-thaw, ensuring that their proliferative capabilities remain intact. © 2025 Wiley Periodicals LLC. Basic Protocol: Isolation, culture, cost-effective expansion, and cryopreservation of caprine satellite cells.

Aptamer-drug conjugates (ApDCs) represent a powerful platform for targeted drug delivery, offering the potential to enhance therapeutic efficacy while minimizing systemic toxicity. However, conventional methods for ApDC preparation often involve complex, multistep procedures with limited control over drug loading and site specificity. This protocol describes a modular and automated strategy for the synthesis and structural optimization of ApDCs using custom-designed phosphoramidites that incorporate anticancer drugs and functional moieties. The customized phosphoramidite is compatible with standard solid-phase oligonucleotide synthesis, enabling the automated and efficient construction of programmable aptamer-drug conjugates (PApDCs) for targeted drug delivery. We also highlight the programmable optimization of aptamer structures to meet clinical needs. The protocol offers a streamlined and versatile platform for the construction of functional ApDCs, facilitating their application in receptor-targeted chemotherapy and nucleic acid-based therapeutics. © 2025 Wiley Periodicals LLC. Basic Protocol 1: Synthesis and purification of phosphoramidites Basic Protocol 2: Automated solid-phase synthesis and characterization of ApDCs.

This protocol presents methods for isolating and identifying novel nucleic acid-binding proteins. We focus on bacterial DNA-binding proteins, although the methods can be readily adapted to identify nucleic acid-binding proteins of eukaryotes or archaea, and proteins that bind to RNAs. Briefly, the DNA sequence of interest is affixed to beads and then incubated with bacterial cytoplasmic extract. Washes with buffers containing nonspecific DNA and low salt concentrations will remove non-adhering and low-specificity DNA-binding proteins, while subsequent washes with higher salt concentrations will elute DNA-binding proteins that have greater specificity. Eluted proteins are then identified by standard proteomic techniques, such as mass spectrometry. © 2025 Wiley Periodicals LLC. Basic Protocol 1: Lysis procedure Basic Protocol 2: DNA affinity chromatography: Borrelia burgdorferi and Leptospira interrogans Basic Protocol 3: Visualization and identification of DNA-binding proteins.

The oncogenic Epstein Barr virus (EBV) is an exclusively human pathogen with related lymphocryptoviruses (γ1‐herpesviruses) only present in monkeys. Therefore, experimentation with EBV infection in a small animal model requires reconstitution or adoptive transfer of human lymphocyte populations, primarily EBV's main host cell, the human B cell. In this protocol we describe human immune system reconstitution after neonatal transfer of CD34+ hematopoietic progenitor cells in lymphodeplete immune compromised mouse strains, using NOD‐scid γc–/– (NSG) mice as a commonly used example. Such reconstituted humanized mice allow intraperitoneal and intranasal infection with EBV and we describe injection of 105 infectious particles of the prototypic EBV strain B95‐8 that can be produced from a recombinant bacmid (p2089) in HEK293 cells. Infection with this dose mimics symptomatic primary EBV infection, infectious mononucleosis (IM), with high viral loads plateauing 4 weeks after infection, with CD8+ T‐cell lymphocytosis at week 5 and 6 after infection. This IM‐like primary EBV infection in humanized mice leads to clonal EBV‐induced B‐cell lymphoproliferations that resemble large B‐cell lymphomas with the latency III program of EBV infection. We describe Basic Protocols to monitor viral loads, immunohistochemistry of infected tissues and spectral flow cytometry to characterize protective T‐cell expansion. The described mouse model has been used by us and others to characterize mutant EBV infections, cell‐mediated immune control of EBV, modulation of EBV pathogenesis by co‐infections with human immunodeficiency virus (HIV) and Kaposi sarcoma associated herpesvirus (KSHV), as well as passive transfer of vaccine elicited antibodies to test their protection against EBV infection. © 2025 The Author(s). Current Protocols published by Wiley Periodicals LLC.Basic Protocol 1: CD34+ human hematopoietic progenitor cell isolation and characterizationBasic Protocol 2: Human immune system reconstitution and characterizationBasic Protocol 3: Recombinant EBV production and humanized mouse infectionBasic Protocol 4: Viral load quantification, lymphoma assessment, and immunohistochemistry after EBV infection of humanized miceBasic Protocol 5: Human T‐cell response analysis after EBV infection of humanized mice