Identification and Ultrastructural Peculiarities of Nestin-Carrying Progenitor Cells in Kidney

TL;DR

This study identifies and examines the unique structure of nestin-positive progenitor cells in kidney cultures using electron microscopy.

Contribution

The study reveals ultrastructural differences between nestin-positive and nestin-negative kidney cells using GFP and electron microscopy.

Findings

Nestin-positive cells showed high ribosome content and protein-synthesizing activity.

Nestin-positive cells had vesicle-containing protrusions and multiple nuclei with high lysosome content.

Tight interactions and structural differences were observed between nestin-positive and -negative cells.

Abstract

In this study, in a culture of renal epithelial cells, we identified those expressing nestin, a cytoskeletal protein associated with stem/progenitor/activated/proliferating cell states. A mouse expressing GFP under the nestin promoter was used, followed by cell isolation and culture. It is hypothesized that this can be used to assess the stem/progenitor/activated/proliferating cell level in a mixed kidney cell culture. Both nestin-positive and nestin-negative cells were demonstrated to be present in the culture. After visualization, cells were attached to a glass slide with a grid, fixed, and prepared for electron microscopy analysis, with each cell visually identified by light microscopy being analyzed. Electron microscopy revealed tight interactions between nestin-positive and nestin-negative cells. Significant differences in the ultrastructure of nestin-positive and nestin-negative cells were observed. Nestin-positive cells were distinguished by a high ribosome content, indicating high protein-synthesizing activity. In the nestin–GFP-high (sorted) population examined by electron microscopy, vesicle-containing protrusions were frequently observed. These cells could contain multiple nuclei of varying sizes and had a high content of lysosomes. No significant differences in mitochondrial ultrastructure were observed in nestin-positive and -negative cells, although functional characteristics evaluated by the membrane potential probe differed.

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1. Introduction

The search for approaches to regenerating tissue damaged by external or internal factors is currently one of the most urgently needed. One solution involves harnessing endogenous reserves, focusing primarily on stem cells, which are found to varying degrees in virtually every organ, including the bone marrow, muscle, heart, brain, blood, skin, liver and kidney. It is believed that, in these organs, stem cells constitute a pool of dormant cells that wake up at the moment when restoration of damaged cells is required after exposure to various factors [1,2,3,4,5]. The pool of stem cells capable of differentiating in the desired direction—replenishing lost, damaged, or underactive specialized cells—is not constant, and its depletion, which occurs over time and depends on the activity of damaging/repairing systems, is a problem that can theoretically be addressed by various approaches [6,7,8,9].

The repair process is crucial for restoring the function of various organs, including the kidney, which can be damaged by physical (e.g., trauma), chemical (exposure to poisons), or biological (bacteria and viruses) factors [10,11]. One of the most common causes is damage resulting from an ischemic attack associated with oxidative stress, which can lead to either a chronic phase requiring extensive pharmacological intervention or the need for hemodialysis, and in the most severe cases, a kidney transplant [12,13,14,15,16]. Neither of these strategies guarantees complete organ recovery, necessitating increased efforts to utilize the body’s internal reserves, particularly the mobilization of resident stem cells [17,18,19,20]. However, this requires accurate identification of stem cells in a biological sample.

The history of the discovery of stem cells dates back to the end of the 19th century; one of the pioneering scientists was Ernst Haeckel [21], who introduced the term Stammzeller (as a derivative of Charles Darwin’s Stambaume, i.e., stem tree). A little later, working on the problem of the development of ascaris, Theodor Boveri used the same term, attributing it to the ability of cells to self-renew and differentiate [22]. An important contribution at the initial stage of the development of stem cell science was made by Valentin Hacker, who in 1882, presented a diagram of the development of the Cyclops embryo, emphasizing the primary cell, which migrates to the center of the embryo and undergoes asymmetric division [23]. In 1907, Artur Pappenheim published a paper in which he placed the cell in the progenitor center of the blood, and may have been the first to introduce the term “stem cell”, which is still used today [24]; others attribute the origin of this term to Alexander Maximov [25].

A particularly important breakthrough occurred in the early 1960s, after which the term “stem cells” ceased to be used for primordial germ cells. This was due in part to the work of Till and McCulloch [26]. As a result, the concept of the presence of primary cells in tissue, capable of proliferating virtually indefinitely and leading to the formation of cells with specialized functions, began to be clearly established. Since then, stem cell science has developed intensively.

Stem cell identification can probably be traced back to the pioneering work describing a protocol for obtaining pluripotent stem cells from mouse embryonic fibroblasts using a cocktail of four transcription factors, later named Yamanaka factors [27]. Currently, the presence of about 25 transcription factors expressed in stem cells has been reported. High levels of expression of these factors in embryonic stem cells, with exceptional suppression in normal somatic cells, have been documented. The Mesenchymal Stromal Cell Committee of the International Society for Cell and Gene Therapy defines mesenchymal stem cells as positive for CD105 (ENG), CD73 (NT5E), and CD90 (THY1), and negative for CD45, CD34, CD14, CD19, and HLA-DR. CD105, CD73, and CD90 are expressed on virtually all mesenchymal stem cells, regardless of the source.

Among the stemness indicators, nestin was suggested to be an important one. It was historically identified in the dentate gyrus of the brain, suggesting the localization of a brainstem niche [28]. However, it was later found in other organs, including the skeletal muscle satellite cells [29], heart [30], testis [30], hair follicles [31], and kidney [32], raising some optimism about the possibility of regulating the regeneration of these vital organs in the event of injury [31,33]. Thus, nestin expression is increasingly viewed as reflecting a progenitor-like or activated cellular state rather than an exclusive marker of undifferentiated stem cells.

So, it remains premature to definitively assert that nestin is a definitive marker of stemness, although most researchers favor this assertion. Nestin is a cytoskeletal protein and a rather unique component of intermediate filaments, characterized by remarkable complexity. Its presence in a variety of tissues and association with various types of normal and cancer cells capable of self-renewal (see review [34]) allows for a rather lenient definition of the association of this protein with progenitor cells. Recent data suggest that it may be more accurate to define the association of nestin with activated, proliferating cells both in normal conditions and during cellular and tissue regeneration [35].

Regardless of the controversial opinions, the presence of nestin in cells is a reporter of a specific cell type possibly involved in regenerative processes, so the identification and characterization of nestin-containing cells is an important research goal. In this study, we used a transgenic mouse expressing GFP under the nestin promoter to identify nestin in different epithelial cell populations. We developed an approach that allows us to compare the structure and ultrastructure of each individual epithelial cell that contains nestin and compare it with the ultrastructure of cells that do not contain nestin.

2. Results

Using confocal microscopy after seeding a dispersed cell suspension on glass-bottom Petri dishes, we obtained images of the fluorescence of GFP-bearing cells and compared them with phase-contrast images of the total cell population on the slide. This revealed a population of cells displaying high levels of GFP fluorescence. It should be noted that not all cells had the same fluorescence intensity, indicating varying levels of nestin expression. Therefore, we selected only cells with high fluorescence intensity (see the criteria of selection in Section 4). Some cells exhibited extremely low fluorescence, likely due to endogenous flavins, and we classified these cells as nestin-negative.

Figure 1 shows an example of such an experiment, in which we photographed cells in the region of interest located in the region of the letter H on a glass grid. We focused on a cell cluster containing a pyramidal, non-fluorescent cell (Figure 1C–E) adjacent to other fluorescent cells. The entire slide containing the cells was fixed, and the region of interest was analyzed by electron microscopy. Individual electron microscopic images were assembled, yielding a complete electron microscopic image of the entire field, comparable to that obtained by conventional microscopy. Figure 1F shows all cells marked in Figure 1A without any gaps.

By focusing on a small group of cells, including the pyramidal-shaped cell, we obtained higher-magnification electron microscopic images to detail the ultrastructure of nestin-positive and nestin-negative cells, which were in close contact with each other. Figure 2 shows that the fluorescent cell exhibits greater osmiophility, resulting in its darker appearance. This occurs due to the increased number of polysomes in nestin-positive cells (see Figures S1 and S4 of Supplementary Material S4), according to which, in nestin-positive cells, the ribosome content was approximately 3 times higher than in nestin-negative cells). This results in a clear distinction between the two cell types separated by a membrane (Figure 2F and Figure S3 of Supplementary Material S4). No clear or pronounced differences in the mitochondrial ultrastructure of the two cells were detected. Figure 1B–D clearly show a population of mitochondria in an orthodox conformation with characteristic cristae, with an abundance of elongated mitochondrial profiles. A similar pattern is observed in fluorescent cells (Figure 2E).

Using the same group of cells, as well as a different cell ensemble, we noted significant differences in plasma membrane morphology between nestin-positive and nestin-negative cells. While nestin-negative cells exhibited the typical structure characteristic of epithelial cells, in which the plasma membrane is typically smooth without outgrowths or structures (Figure 3A,B), the overwhelming majority of nestin-positive cells had a bumpy surface with outgrowths. Furthermore, nestin-positive cells could display multiple nuclear profiles (see Figure 3A), indicating either the presence of multiple nuclei or an extremely uneven nuclear surface.

In another example, fluorescent and non-fluorescent cells were also selected, but they did not interact with each other to exclude direct influence from their partners. In this case, clear differences in ribosome content (see Supplementary Material S4, Figure S3) and ultrastructure (Figure 2, Figure 3 and Figure 4) were also observed, expressed in the very frequent formation of vesicular structures or in the attachment of containers filled with these structures.

Our analysis showed that observed nestin-positive cells have the ability to form vesicular structures, or containers, in which these structures are placed. Here, we use the term “container” to describe membrane-bound protrusive structures containing multiple vesicular structures, based on their consistent ultrastructural appearance. Figure 5 shows examples that may correspond to different stages of vesicle container rejection from the cell. Furthermore, a characteristic and often observed feature of nestin-positive cells was the presence of a large number of lysosomes, sometimes containing mitochondrial structures (Figure 5B), which was noted in the electron microscopy images.

We did not observe a statistically significant difference in the quantitative characteristics of mitochondrial size in the two cell types (see Supplementary Material S1); however, judging by the TMRE (tetramethylrhodamine ethyl ester) fluorescence intensity, the overwhelming majority of nestin-negative cells had a lower membrane potential. This is evident in the different analyses shown in Figure 6B–D.

To exclude the possibility that nestin-negative cells represent quiescent cells, given that nestin expression is cell cycle–dependent, we compared two cell culture conditions containing nestin-positive and nestin-negative cells. These conditions were time-dependent when comparing the second and third days (2 DIV and 3 DIV, see Supplementary Material S3) of dispersed kidney tubule culture. By the third day of culture, an almost complete cellular monolayer is formed, and this occurs due to both an increase in GFP-negative cells (see Supplementary Material S3) and GFP-positive cells (see Figure 3 in [8]). This shows that GFP-negative cells also divide, and the division occurs quite intensively.

It is also unlikely that the observed differences in phenotype are due to cell culturing. Analysis of kidney sections from nestin mice (Supplementary Material S2) clearly shows GFP fluorescence exclusively in a number of renal tubules, and these are the source material for the mixed culture that was studied after 2–3 days of culturing.

3. Discussion

According to our data, the task we set—namely, the assessment of the cell ultrastructure in a selected individual cell possessing specific properties—has not been undertaken by anyone due to the high labor costs and complexity of implementation. In 1988, we implemented such a task using human fibroblasts or neonatal rat cardiomyocytes as objects [36], and an individual mitochondrion was damaged by a focused laser beam. After this damage, the cell was fixed, and a series of ultrathin sections were analyzed by electron microscopy to identify the damaged locus and changes in mitochondrial conformation using a three-dimensional reconstruction of the host cell.

In this study, we reproduced this approach with the sole purpose of identifying a nestin-producing cell in cell culture and assessing the ultrastructure of this specific cell. We also compared nestin-positive and nestin-negative cells based on their ultrastructural features.

We attributed nestin-positive cells to cells with stemness or/and progenitor or/and proliferative regenerative characteristics, which, according to our data and those of other researchers, distinguishes such cells from differentiated cells.

With our extensive expertise in mitochondrial science, we have largely focused in the past on the behavior of mitochondria in stem cells. One of the most striking properties of stem cells (and cancer cells as well) has been their ability to donate mitochondria to other cells experiencing various problems. Mitochondria are transferred from stem cells to differentiated cells either through tunneling nanotubes [37,38,39,40,41], through gap junctions [42,43,44], or through direct release of mitochondria into the extracellular environment [45,46,47,48,49].

We found no significant differences in the mitochondrial ultrastructure of nestin-positive and nestin-negative cells. This does not preclude the need for a more detailed and exhaustive analysis of cristae configuration; the extent of their local and global changes; the density of the mitochondrial matrix, which largely reflects the degree of mitochondrial coupling; the integrity of the outer mitochondrial membrane; local swelling; and contacts with other intracellular elements. Such an analysis not only requires extensive statistical data but also the mandatory use of automated analysis, which is currently imperfect. After analyzing the mitochondrial morphology (length, thickness, and area occupied by mitochondria in the two cell types), we did not find any significant differences (see the Supplementary Material S1); however, after segmentation of mitochondria in the cell and assessing the intensity of TMRE staining—which reports on the magnitude of the transmembrane potential on the inner membrane of the mitochondria—some difference was revealed, indicating a modest but significant difference in the functioning of mitochondria in the two cell types.

The difference in the transmembrane potential on the inner mitochondrial membrane between nestin+ and nestin- cells appears small, but to assess the influence of membrane potential on intracellular processes, it is important to understand that large changes associated with normal physiology are generally not expected. The range over which changes in mitochondrial membrane potential occur under physiological loads is only a few tens of millivolts. While in isolated mitochondria, the membrane potential is in the range of 180–200 mV in a state of complete rest, which is called state 4 [50,51,52], with full activation of mitochondrial respiration (by adding ADP or an uncoupler), called state 3, it decreases to 150–180 mV. Therefore, it is in this 20–30 mV interval that all physiologically acceptable changes occur [53]. Of course, in mitochondria, there are also values that go beyond this interval (values above 220 mV, which is called hyperpolarization, and values below 120–150 mV up to the complete disappearance of the membrane potential (depolarization) during the generation of a mitochondrial transition pore in mitochondria (megachannel)) [54,55,56]. However, both of these conditions, if prolonged, are signs of pathology, often signaling the possibility of a lethal cascade [57,58]. Thus, normally, and when comparing absolute changes in membrane potential, the values in different cells may be small and not easily detectable, although they may have important metabolic significance.

To understand how significant small changes in membrane potential are, one must understand the importance of membrane potential for cellular functioning. The membrane potential of mitochondria (minus inside) is generated and determined by the activity of three proton pumps (complexes I, II, and IV). It is consumed by the rotational movement of the ATP synthase complex motor, thereby lowering the membrane potential. Proton pumps accordingly adapt to the activity of ATP-consuming systems, which has been formulated as a “push” and “pull” mechanism [59]. As a result, in a mitochondrion in a resting state, the ADP levels are low and ATP is high, which corresponds to a state of respiratory control where mitochondrial respiration is minimal, determined only by the ion leak through the inner membrane [60]. In the activated state (ATPase activity in the cell is high), ADP in the cell increases, respiration is activated, and the membrane potential decreases slightly. This drop in the potential has a lower limit at state 3 (or state 3u in the presence of an uncoupler), when the respiration rate is not limited by the availability of substrates or the activity of the ATP/ADP translocator [61,62], but is limited only by the activity of the respiratory chain.

The importance of membrane potential in the cell is not limited to driving the rotation of the components of the ATP synthase complex [63]. It is critical for providing the transport of proteins [64] and cations [65] into the mitochondria, nucleotide metabolism in the mitochondria [66], participation in the mitochondrial quality control mechanism [67], initiation of potassium energy [68,69,70], etc. [63]. The fact that under conditions of an energy crisis caused by the onset of hypoxia, mitochondria unable to synthetize ATP use cellular ATP by reversing the ATP synthase reaction, creating a membrane potential [71], speaks of the crucial role of the mitochondrial membrane potential for the functioning of mitochondria, while homeostasis of the membrane potential is one of the mandatory prerequisites for the normal existence of mitochondria [63].

It should be noted that nestin-positive cells formed tight contacts with nestin-negative cells in culture, and the border between them was not always easy to identify. However, nestin-positive cells differed from nestin-negative cells—even those with which they were in contact—by a higher ribosome content (see Figures S2 and S3 of Supplementary Material S4), which gave these cells a darker staining in the electron microscopy image (higher osmiophility). This indicates a trend to higher protein-synthesizing activity in nestin-positive cells. Nestin-positive cells very often exhibited a very high content of lysosomal structures of varying configurations, indicating active autophagy and mitophagy, i.e., a high level of cellular turnover.

Another striking difference was the organization of the plasma membrane. While nestin-negative epithelial cells had a relatively smooth surface, nestin-positive plasma membranes formed protrusions, often culminating in the obvious generation of extracellular vesicular structures. Although the nature of these vesicular structures was not directly assessed in this study, their morphology and dynamics are consistent with previously described extracellular vesicle–producing phenotypes of stem and progenitor cells. This is consistent with the paradigm of the massive generation of extracellular vesicles by stem cells [72,73]. All this points to the active participation of stem cells in intercellular communication, where they largely act as donors of their intracellular components, ensuring a three-way change in metabolism in acceptor cells through killing, healing, and rejuvenation [74,75].

The high membrane potential in nestin-positive cells may have various causes and consequences. If we consider the aforementioned “push” and “pull” mechanism [59], the second part of this mechanism (“pull”) reflects passive leak through the bilayer, or through proteins specializing in creating regulated leak (such as uncoupling proteins), as well as by increasing the potential due to intracellular endergonic reactions (primarily those associated with ATP hydrolysis). Since the electron microscopy images demonstrate a high content of ribosomes, this suggests high protein-synthesizing activity in nestin-positive cells. At the same time, these cells exhibit high levels of lysosomal activity and the active formation and release of exosomes from the cell, which, like protein synthesis, are very energy-consuming processes. All of these processes, which discharge membrane potential, are the main components of the “pull” mechanism. From all this, we can speculate that a higher potential is determined by very high-energy-producing activity, exceeding consumption. This may reflect higher activity of proton pumps (“push” component) associated with low leak, which will result in lower oxygen consumption (closer to the respiratory control state). Overall, this reflects higher metabolic activity of cells producing nestin, which is presumably an indicator of regeneration. Given the direct relationship between membrane potential and redox potential, the redox potential in nestin+ cells should be in a more reduced state, which gives the cell greater resistance to oxidative stress.

Taken together, our data propose that nestin-positive cells in renal epithelial cultures represent a distinct ultrastructural and functional phenotype characterized by high biosynthetic activity, active vesicle formation, and increased lysosomal turnover. The lack of major differences in mitochondrial architecture, along with subtle functional changes, suggests that metabolic adaptation accompanies rather than drives this phenotype. Importantly, the correlative approach used allows direct linkage of light-microscopic identity with ultrastructural and functional features at the level of individual cells.

4. Materials and Methods

4.1. Mice

The study was performed on the male nestin–GFP transgenic reporter mouse strain [76]. The nestin–GFP mice were generated [77] and kindly provided by Grigori Enikolopov. The experiments were carried out on homozygous transgenic mice, 21 days old. Animal protocols were evaluated and approved by the Animal Ethics Committee of the A.N. Belozersky Institute of Physico-Chemical Biology, Lomonosov Moscow State University, Protocol 3/19 from 18 March 2019. All procedures were in accordance with the “Animal Research: Reporting of In Vivo Experiments” (ARRIVE) guidelines. The animals had unlimited access to food and water and were kept in cages in a temperature-controlled environment (20 ± 1 °C) under a 12/12 h light/dark regime.

4.2. Culture of Mouse Renal Tubular Cells

Primary cultures were isolated from the kidneys of nestin–GFP mice. The mice were euthanized, and kidneys were aseptically isolated, cut into small pieces, and incubated with 0.25% collagenase II type (Gibco, ThermoFisher Scientific, Waltham, MA, USA) in DMEM/F12 bicarbonate-free media at 37 °C for 15 min. Kidney pieces were pipetted, and the resulting suspension was centrifuged for 5 min at 400× g to pellet the tubular fraction. The pellet was resuspended in a complete DMEM/F12 culture medium with 10% fetal bovine serum (FBS). The resulting renal tubules were plated on Mattek culture dishes with grid. After 48 h, the medium was changed to remove cellular debris. After 3–4 days of cultivation, the cells reached the monolayer and were analyzed on a confocal microscope, followed by fixation for electron microscopy.

4.3. Imaging

Cellular GFP fluorescence and transmitted light were imaged using confocal and conventional fluorescent microscopy. GFP fluorescence was evaluated using a 488 nm excitation wavelength with emission collected at 500–530 nm, correspondingly using an LSM 900 confocal microscope (Carl Zeiss, Jena, Germany) or Nikon Eclipse Ti (Nikon, Tokyo, Japan). Under our experimental conditions, the possibility that mature renal cells are GFP-positive due to the different turnover of GFP and nestin is quite small, since the half-life of GFP in the cells is about 26 h [78]; thus, we classified all cells with GFP fluorescence as nestin-expressing cells. For the selection criterion, we used the histogram of GFP fluorescence distribution for the entire sample; cells with a fluorescence intensity of less than 5% of the maximum in the histogram were considered nestin-negative, and all other cells with a fluorescence intensity higher than 50% of the maximum were considered nestin-positive.

4.4. Segmentation of Mitochondria

Segmentation of mitochondria in tetramethylrhodamine ethyl ester (TMRE, 200 nM, 30 min incubation) stained nestin-negative and nestin-positive renal cells. Twenty-five different fields with cells were analyzed. The fields contained different numbers of cells: a minimum of 1–2 single cells in some fields, and a maximum of about a dozen cells forming islets in other fields. The total number of cells analyzed was just slightly over 100 (the borders between cells are not always easy to determine, even with an electron microscope). The area occupied by mitochondria and the membrane potential evaluated by TMRE staining in both types of cells was recorded after mitochondria segmentation. Snapshots of mitochondria in the red channels were saved in the “tiff” format. Frames in the red channel were processed by the MitoSegNet neural network model [79] to obtain mitochondrial segmentation masks (basic launch parameters and a filter for the minimum object size of 30 pixels were used). Morphological and functional characteristics of individual mitochondria were analyzed using the scikit-image Python library (version 0.26.0) [80]. Statistical analysis was performed using the statannotations Python library [81]. Comparisons between groups were made by a Mann–Whitney U test, and p-value ˂ 0.05 was considered statistically significant.

4.5. Electron Microscopy

For transmission electron microscopy, cells seeded onto glass-bottom Petri dishes with grid were used. After the desired cells were imaged on a confocal microscope and localized in the grid, the cells were fixed in a mixture of 2.5% glutaraldehyde–2% paraformaldehyde in 0.1M sodium cacodylate buffer, pH 7.2, for 12–24 h, washed in buffer, and post-fixed in 1% osmium tetroxide for 1 h at 4 °C in the same buffer. The samples were then dehydrated in an ascending series of ethanols, transferred to propylene oxide, and infiltrated in EMbed 812 resin (Electron Microscopy Sciences, Hatfield, PA, USA) using the sequence: propylene oxide:resin 2:1, 1:1, and 1:2 throughout 24 h (8 h each), and pure resin for 24 h kept for 48 h at 65 °C for polymerization. The glass from the Petri dish was removed by a series of transfers from liquid nitrogen to hot water and back. Areas of interest containing specific cells imaged by confocal microscopy were marked, selected, and sectioned in Leica ultramicrotomes (Leica Microsystems, Vienna, Austria). Visualization of the cells was performed using a JEM1400 electron microscope (JEOL, Tokyo, Japan) equipped with a QUEMESA camera (Olympus, Center Valley, PA, USA). Thin sections (40–70 nm thick) were mounted on nickel grids and stained in uranyl acetate and lead citrate.