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The Ameliorative Effect of Dexamethasone on the Development of Autoimmune Lung Injury and Mediastinal Fat-Associated Lymphoid Clusters in an Autoimmune Disease Mouse Model

In our previous study, we revealed the ameliorative therapeutic effect of dexamethasone (Dex) for Lupus nephritis lesions in the MRL/MpJ-Fas lpr/lpr (Lpr) mouse model. The female Lpr mice developed a greater number of mediastinal fat-associated lymphoid clusters (MFALCs) and inflammatory lung lesions compared to the male mice. However, the effect of Dex, an immunosuppressive drug, on both lung lesions and the development of MFALCs in Lpr mice has not been identified yet. Therefore, in this study, we compared the development of lung lesions and MFALCs in female Lpr mice that received either saline (saline group “SG”) or dexamethasone (dexamethasone group “DG”) in drinking water as a daily dose along with weekly intraperitoneal injections for 10 weeks. Compared to the SG group, the DG group showed a significant reduction in the levels of serum anti-dsDNA antibodies, the size of MFALCs, the degree of lung injury, the area of high endothelial venules (HEVs), and the number of proliferating and immune cells in both MFALCs and the lungs. A significant positive correlation was observed between the size of MFALCs and the cellular aggregation in the lungs of Lpr mice. Therefore, this study confirmed the ameliorative effect of Dex on the development of lung injury and MFALCs via their regressive effect on both immune cells’ proliferative activity and the development of HEVs. Furthermore, the reprogramming of MFALCs by targeting immune cells and HEVs may provide a therapeutic strategy for autoimmune-disease-associated lung injury.

Introduction
The term autoimmunity represents a condition wherein the body's natural immune defense system cannot differentiate between its own healthy tissues and potentially harmful or foreign antigens. This causes the body to mistakenly generate an immune response against its own host cells [1]. More than 80 types of autoimmune diseases have been reported in humans, and they are found to affect different body organs, including rheumatoid arthritis (RA) in the joints [2]; inflammatory bowel arthritis (Crohn's disease and ulcerative colitis) in the digestive tract [3]; dermatomyositis in the skin and muscles [4]; and systemic lupus erythematosus (SLE) in the joints, kidneys, heart, and skin [5]. Interestingly, pulmonary involvement, including interstitial lung disease and interstitial fibrosis, has also been reported in many autoimmune diseases, such as systemic sclerosis, SLE, and RA. These are referred to as autoimmune lung diseases [6].
Autoimmune diseases have been reported to affect both human beings (8% of the population) [7] and domestic animal species [8]. Several murine autoimmune mouse models have been successfully used to study the pathogenicity of the disease in humans. The MRL/MpJ-Fas lpr/lpr (Lpr) mice are one of the autoimmune disease mouse models used frequently as an SLE model. They are widely used because they exhibit lesions that closely mimic the lesions found in humans. This includes lymphoproliferation, splenomegaly, lymphadenopathy, and injured lungs and kidneys. The autoimmunity is due to the increased titer of anti-double-strand DNA (dsDNA) autoantibodies in the blood [9,10]. Furthermore, we previously demonstrated a significant positive correlation between the development of lung injury and the size of the novel immunological cluster associated with mediastinal adipose tissue, which was termed mediastinal fat-associated lymphoid clusters "MFALCs" in Lpr mice [11].
Leukocyte recruitment in the lung plays a major role in the pathogenesis of lung injury. Such recruitment mainly depends on the function of chemokines and their receptors [12]. Interestingly, both chemokine (C-X-C motif) ligand 9 (CXCL9) and CXCL13 are linked to the development of autoimmune inflammatory disorders, particularly during incidences of lung injury [13,14].
Dexamethasone is one of the corticosteroids that exhibits anti-inflammatory and immunosuppressant effects [15]. Recently, the therapeutic impact of dexamethasone on the degree of lung damage associated with COVID-19 has also been reported [16]. Additionally, we have previously revealed its therapeutic effect on lupus nephritis in the Lpr autoimmune disease mouse model [17]. However, the effect of dexamethasone on the degree of lung injury and the development of MFALCs in Lpr mice remains undefined.
In the prevalence of autoimmune diseases, sex-related differences are reported in human patients, with more incidence observed in females [7]. Additionally, our previous report revealed increased development of MFALCs and lung injury in female Lpr mice compared to that of male mice [18]. In this study, we investigated the effect of dexamethasone on the lymphoproliferation of MFALCs and spontaneous immune cell infiltration within the lungs of female Lpr mice. Our results highlighted the potential ameliorative effects of dexamethasone on the development of lung injury and MFALCs. Additionally, we suggest that the underlying mechanism of such therapeutic impact may be via their regressive effect on both the immune cells' proliferative activity and the development of high endothelial venules (HEVs), along with the suppression of chemokines. Our previous reports revealed the role of MFALCs in the development of lung injury in the Lpr autoimmune disease mice model [11,18]. However, the impact of immunosuppressive drugs on both lung lesions and the development of MFALCs in Lpr mice has not been identified yet. Therefore, in the current study, we examined sections of lung and mediastinal fat tissues (MFTs) stained with hematoxylin and eosin (HE) in both saline and dexamethasone groups. The lung tissue in the saline group showed thickening of the interalveolar septa and numerous peribronchial and perivascular mononuclear cell aggregations (MNCA) that replaced normal lung architecture with numerous collapsed alveoli. In contrast, the lung tissue of the dexamethasone group mice showed alleviation in the degree of MNCA, indicating a restoration of normal lung structure ( Figure 1A). Interestingly, compared to the saline group, the morphometric measurement in the dexamethasone group revealed a significant decrease in the ratio of the area of MNCA/area of the lung field, suggesting ameliorative effects of dexamethasone on lung injury ( Figure 1C). MFALCs (blue arrows) associated with the MFTs. (C,D) Graphs indicating the morphometrical data of the percentages for the ratios of the area of MNCA/area of the lung field (mm 2 ) and the area of LCs/the total area of MFTs (mm 2 ). (E) Graph indicating the serum levels of anti-doublestranded DNA autoantibodies. Highly significant values (**) were observed between the saline and dexamethasone groups (p < 0.01), where n = 5 in each experimental group. Analysis was conducted using the Mann-Whitney U test. Data are presented as mean values ± SE.

Results
Simultaneously, we investigated the MFTs in both study groups. Notably, the saline group showed numerous and prominent lymphoid clusters (LCs) within the MFTs, which were associated with few adipose tissues. Controversially, the dexamethasone group showed fewer and smaller size LCs but with the association of numerous adipose tissues ( Figure 1B). Similarly, we also performed a histoplanimetrical analysis, where the ratio of the area of LCs/total area of MFTs was used as an index to determine the degree of the development of MFALCs between the saline and dexamethasone groups. Remarkably, a significant decrease was observed in the percentage of this ratio in the latter group compared to the former ( Figure 1D).

The Degree of Systemic Autoimmunity in the Saline and Dexamethasone Groups
Degree of lung injury and MFALC development is significantly associated with systemic autoimmunity [18]. Therefore, to examine the impact of dexamethasone treatment on systemic autoimmunity, we evaluated the titer of anti-double-stranded DNA autoantibodies (anti-dsDNA abs) in the sera of mice from both the saline and dexamethasone groups. Intriguingly, the dexamethasone group showed a significantly lower level of anti-dsDNA abs compared to that of the saline group ( Figure 1E).

Immune Cells in the Lung and MFALCs of Saline and Dexamethasone Groups
Previously, we characterized the immune cell populations in the lung and MFALCs of Lpr mice and revealed that both lymphocytes and macrophages are predominant populations [18]. To further investigate the impact of dexamethasone on immune cell populations in both the lungs and MFALCs, the morphometrical changes were compared among the studied groups. As shown in Figure 3 and Figure S1, immunohistochemical staining and morphometrical analysis of the positive index ratios (the positive area of the immune cells/field area) were used to compare the composition of immune cells (CD3 + T-lymphocytes, B220 + B-lymphocytes, Iba1 + macrophages, and Gr1 + granulocytes) in the lungs and MFALCs between the saline and dexamethasone groups. Except for the lung granulocytes, the dexamethasone group showed decreased values for the percentages of positive immune cell index ratios in both the lungs and MFALCs compared to the saline group. No significant difference was observed in the percentages of T-lymphocytes positive index ratios in either the lungs or MFALCs between the study groups (CD3 + T-lymphocytes, Supplementary Figure S1A). However, a highly significant decrease was observed in the B-lymphocyte values in the dexamethasone group compared to the saline group (B220 + B-lymphocytes, Supplementary Figure S1B). In the case of macrophage positive area ratios, non-significant and significant differences were observed in the lungs and MFALCs, respectively (Iba-1 + macrophages, Supplementary Figure S1C). Moreover, for granulocyte positive area ratios, a highly significant increase and decrease were observed in the lungs and MFALCs, respectively (Gr-1 + granulocytes, Supplementary Figure S1D).
To characterize the Iba-1 + macrophage subsets in the lung tissue, triple immunofluorescence staining was performed with Iba-1 (macrophage marker), CD68 (M1 macrophage marker), and CD204 (M2 macrophage marker) antibodies. Interestingly, the examination of immunofluorescent-stained lung sections showed a decrease in the Iba-1 + macrophages and CD68 + M1 macrophages and an increased expression of the CD204 + M2 macrophages in the dexamethasone group compared to that of the saline group (Supplementary Figure S2).

Analysis of the Degree of Proliferation of Immune Cells in the Lungs and MFALCs between the Saline and Dexamethasone Groups
Immunohistochemical staining of Ki67 (a proliferation marker) was performed in the lungs and MFTs of both study groups to evaluate the degree of proliferation of immune cells, which is a major characteristic of Lpr mice. Compared to the saline group, the dexamethasone group showed fewer Ki67 + proliferating cells in the lungs ( Figure 2A) and MFALCs ( Figure 2C). Consistent with our histological observations, the morphometrical analysis revealed a highly significant reduction in the percentages of the ratio of Ki67 + proliferating cell density (number/field area) in the lungs ( Figure 2B) and the ratio of Ki67 + proliferating cell number/total immune cell numbers in the MFALCs ( Figure 2D) in the dexamethasone group compared to the saline group. To address the proliferating immune cell types, dual immunofluorescent staining of the lung sections was performed. The lung sections of both studied groups were co-stained with the anti-Ki67 antibody along with immune cell markers (anti-B220 Supplementary Figure S3, anti-Iba1 Supplementary Figure S4, and anti-Gr1 Supplementary Figure S5 antibodies). Interestingly, high and low numbers of Ki67 + B220 + B-lymphocytes were observed in the lungs of the saline and dexamethasone groups, respectively (Supplementary Figure S3). However, few Ki67 + Iba1 + macrophages were observed in the lungs of both studied groups (Supplementary Figure S4). On the other hand, low and high numbers of Ki67 + Gr1 + granulocytes were observed in the lungs of the saline and dexamethasone groups, respectively (Supplementary Figure S5).

Comparison of HEVs and the Development of Lymphatic Vessels (LVs) in the Lung and MFALCs between the Saline and Dexamethasone Groups
Our previous reports revealed the role of both HEVs and LVs in the development of MFALCs and lung injury [19,20]. To investigate the degree of HEVs and the development of LVs in the lungs and MFTs, immunostaining was performed using peripheral node addressin (PNAd) and lymphatic vessel endothelial hyaluronic acid receptor-1 (LYVE-1) antibodies, respectively. Interestingly, the dexamethasone group showed fewer prominent HEVs in the lungs ( Figure 4A) and MFALCs ( Figure 4B) compared to those of the saline group. Next, we performed a morphometrical analysis of the percentage of HEV relative area ratios (area of the HEVs/area of the field) in the lungs ( Figure 4C) and MFALCs ( Figure 4D). Importantly, statistical analysis revealed a highly significant reduction in this ratio in the dexamethasone group compared to that of the saline group in both lungs and MFALCs ( Figure 4C,D). For the examination of the degree of LV development in the lung, dual immunofluorescent staining for LYVE-1/CD68 was performed. Interestingly, more numerous LYVE-1 + LVs and CD68 + macrophages were observed in the lung sections of the saline group than in those of the dexamethasone group. Furthermore, most of the LVs in the lungs of the saline group, but not of the dexamethasone group, were engorged with immune cells (Figure 5A), and their lining epithelial showed a positive reaction to Ki67 immunostaining (Supplementary Figure S3). Simultaneously, morphometrical analysis of the LYVE-1 + LVs relative area ratio was performed for the LYVE-1 immunohistochemical-stained lung and MFALC sections. The dexamethasone group mice showed a highly significant reduction in this ratio in both lungs and MFALCs in comparison to that of the saline group mice ( Figure 5B

Analysis of the Impact of Dexamethasone on Chemokine Expression in the Lungs between the Saline and Dexamethasone Groups
We examined the expression of CXCL9, CXCR5, and CXCL13 in the lungs of both study groups. Interestingly, CXCL9 was previously reported to induce the chemotaxis, differentiation, and proliferation of leukocytes, especially CD8 and CD4 positive T lymphocytes [21,22]. Therefore, double immunofluorescence was performed to detect CXCL9 + cells along with either CD4 + or CD8 + T lymphocytes in the lung sections of both study groups. As shown in Supplementary Figure S6, numerous CXCL9 + , CD4 + , and CD8 + cells were observed in the lungs of the saline group compared to those of the dexamethasone group. Of note, many CXCL9 + cells were mainly observed within the area of MNCA in the lungs of the saline group mice. Moreover, numerous CXCL9 + cells showed co-staining with the CD8 + T lymphocytes (Supplementary Figure S6B) but not with CD4 + T lymphocytes (Supplementary Figure S6A) in the lungs of the saline group rather than the dexamethasone group.
Importantly, the CXCR5 receptor and its ligand (CXCL13) have been reported to be involved in B cell-selective chemotaxis and recruitment [23,24]. Thus, the co-expression of either CXCR5/CD79a positive B lymphocytes or CXCL13 and B220 positive B lymphocytes was examined in the lungs of both study groups. As shown in Figure 6A, more positive CXCR5 and CD79a cells were observed in the lungs of the saline group than in those of the dexamethasone group. Interestingly, the lungs of the saline group, but not the dexamethasone group, revealed numerous CXCR5 +/ CD79a + B lymphocytes co-stained cells. Additionally, more positive CXCL13 and B220 cells were observed in the saline group compared to the dexamethasone group. In the saline group, numerous B220 + B-lymphocytes were mainly observed within the MNCA. However, the CXCL13 + cells were mainly observed within the vicinity of the MNCA ( Figure 6B). Furthermore, a significantly higher percentage for the ratio of CXCR5 + and CXCL13 + cell densities (number/mm 2 lung area) was observed in the saline group compared to that of the dexamethasone group ( Figure 6C,D). Representative merged images of double immunofluorescent staining for CXCR5 "red" and CD79a "green" positive cells, and Hoechst nuclear staining "blue" (A) along with CXCL13 "green" and B220 "blue" positive cells, and Hoechst nuclear staining "white" (B). Notice positive reactions for CXCR5 (red arrows), B-lymphocytes (blue arrows), CXCL13 (violet arrows), and CXCR5 +/ CD79a + co-stained cells (yellow arrows). (C,D) Representative graphs showing the percentage for the ratio of positive cell density for the CXCR5 (C) and CXCL13 chemokines (D). A highly significant value (**) was observed between saline and dexamethasone groups (p values < 0.01), where n = 5 in each experimental group. Analysis was conducted using the Mann-Whitney U test. Data are presented as mean values ± SE. Figure 7A,B shows significant positive correlations among the size of MFALCs, the lung MNCA area ratio ( Figure 7A), and the HEV area ratio in the MFALCs ( Figure 7B). Interestingly, the correlations among the HEV area ratio, MFALC HEVs ( Figure 7C), and MNCA in the lungs ( Figure 7D) also show a highly significant positive correlation ( Figure 6C,D). This suggests the possible role of HEVs in the chemoattraction of the immune cells of MFALCs to the lungs and also the development of numerous MNCA in the lungs of the saline group mice.

Analysis of Histopathological Correlations between Immune Cell Populations and the Proliferating of Cells in the Lungs and MFALCs of Both Saline and Dexamethasone Group Mice
The correlations between the immune cell populations (B220 + B-lymphocytes, CD3 + T-lymphocytes, Iba1 + macrophages, and Gr-1 + granulocytes) and the degree of proliferation of the cells in the lungs and MFALCs were analyzed in both the study groups ( Figure 8). Interestingly, the B-lymphocyte population in the lungs showed a highly significant positive correlation with MFALCs ( Figure 8A). Furthermore, a significant and positive correlation was observed in the proliferation of cells among the lungs and MFALCs of both the study groups ( Figure 8E). However, the T-lymphocyte ( Figure 8B) and macrophage ( Figure 8C) populations showed a non-significant positive correlation between the lungs and MFALCs, while a non-significant negative correlation was observed between the lungs and MFALCs granulocytes ( Figure 8D).

Discussion
The present study examined the impact of dexamethasone, an anti-inflammatory corticosteroid, on the degree of lung injury and the development of MFALCs in the Lpr autoimmune mouse model. This mouse model was previously reported to develop severe lung injury and prominent MFALCs associated with autoimmunity [11]. Corticosteroids, especially dexamethasone, have been successfully used in the treatment of patients with acute lung injury, COVID-19, and idiopathic pulmonary fibrosis [25][26][27]. Additionally, our recent study revealed that the administration of dexamethasone in the Lupus nephritis Lpr autoimmune mice model led to the alleviation of renal lesions via the reduction in vasculature-associated lymphoid tissues in the kidney of Lpr mice [17]. However, no clarifications are available on the effect of dexamethasone on the degree of lung injury and the development of MFALCs. Moreover, the correlations between MFALC development and the amelioration of lung injury after the administration of dexamethasone have not been clarified yet. Therefore, we aimed to clarify the possible mechanisms through which dexamethasone exerts its ameliorative effect. We examined the lungs and MFTs of both saline-and dexamethasone-administered Lpr mice. Interestingly, our previous study revealed sex-related differences among the Lpr mice in the presentation of the degree of lung injury and development of MFALCs, with the number of lung lesions and prominent MFALCs being greater in female mice [18]. Furthermore, women are reported to be more susceptible to autoimmune diseases compared to men [1]. Therefore, instead of male mice, we used female Lpr mice in this study.
Our results showed that, compared to the saline group, the lungs of the dexamethasone group mice showed less MNCA along with the restoration of normal alveolar morphology, which suggests an ameliorative effect of dexamethasone on the severity of lung injury. This was also in line with the results presented by Villar et al. [28], who reported the efficacy of dexamethasone in the treatment of acute respiratory distress syndrome in patients with COVID-19. Next, we also confirmed that the dexamethasone-administered group showed a significant decrease in the size and number of MFALCs. Our results revealed a significant positive correlation between the degree of lung injury and the development of MFALCs, which strongly suggested a major role for MFALCs in the development of lung injury. Notably, the dexamethasone group showed a significant decrease in the serum anti-dsDNA abs (as an autoimmune index) compared to that of the saline group. Therefore, we postulated that the administration of dexamethasone could control the development of MFALCs and lung injury via a systemic effect.
Also, HEVs and LVs play a major role in the lymphoid organs and tissues. HEVs are specialized postcapillary venules composed of cuboidal blood endothelial cells. These play a role in chemoattracting naïve and central memory cells. Furthermore, the role of LVs has been reported to include transporting antigens, lymphocytes, and antigen-presenting cells in and out of the lymph nodes [29]. Importantly, our previous study revealed a major role for both HEVs and LVs in the development of both lung injury and MFALCs in several mice models [11,19,20]. Predictably, our present investigation revealed less developed HEVs and LVs in both the lungs and MFALCs of the dexamethasone-administered group compared to those of the saline group. Moreover, significant positive correlations were observed between the MNCA and the lung HEVs, as well as between the size of MFALCs and their HEVs. Additionally, since our results revealed a positive and highly significant correlation between the development of HEVs both in the lungs and MFALCs, we strongly suggest a major role for both HEVs and LVs in the development of lung injury and MFALCs. Additionally, we suggest that the ameliorative effect of dexamethasone may be induced via its interference with the development of both HEVs and LVs.
To understand the impact of dexamethasone on the intrathoracic immune hemostasis, we examined the immune cells and their degree of proliferation in both the lungs and MFALCs of dexamethasone and saline group mice. Our results revealed that, compared to the saline group, all the studied immune cell populations were alleviated, except for the lung granulocytes, in the lungs and MFALCs of the dexamethasone-administered mice. A non-significant difference was observed in the alleviation of T-lymphocytes in the lungs and in macrophages and T-lymphocyte populations within the MFALCs. However, a highly significant reduction was observed in the B-lymphocyte populations within the lungs and MFALCs of the dexamethasone group mice. Additionally, compared to the saline group, a significant reduction was observed in the CXCL13 chemokine within the lungs of the dexamethasone group mice. CXCL13 has been reported to play a major role in the pathogenesis of several autoimmune diseases, lymphoproliferative disorders, inflammatory conditions, as well as lymphoid neogenesis [30]. Moreover, this chemokine is known as B cell-attracting chemokine 1 or B lymphocyte chemoattractant, and it plays an important role in the process of selectively attracting B lymphocytes [31]. Interestingly, the lungs of the saline group mice revealed several CXCL13 + cells in the confines of MNCA that showed numerous B-lymphocyte populations. Therefore, we postulated that dexamethasone could induce alleviation in the B-lymphocyte population via the suppression of CXCL13. Furthermore, the lungs of the saline group revealed a significantly higher number of CXCR5 + cells as well as more numerous CXCR5 + B220 + B lymphocytes, especially within the MNCA, compared to the dexamethasone group. Interestingly, recent reports have focused on the role of CXCL13 and its receptor "CXCR5" as a B-cell chemoattractant, especially in autoimmune diseases [30].
The current investigation also examined the role of CXCL9 in the recruitment of CD4 + helper and CD8 + cytotoxic T lymphocytes in the lung tissue of the studied groups. More expressions for CXCL9, CD4, and CD8 were observed in the saline group lungs than those of the dexamethasone group. Furthermore, our results revealed the co-expression of CXCL9 along with CD8 + cytotoxic T lymphocytes in the lungs of the saline group in contrast with the dexamethasone group. Similarly, the role of CXCL9 in CD8 + cytotoxic T lymphocytes has been recently clarified [32].
Several reports have clarified the role of macrophage polarization in the pathogenesis and resolution of lung injury [33]. There are two main macrophage subsets that are involved in the pathogenesis of inflammation progression or recovery: the classically activated (or inflammatory) M1 macrophages and the alternatively activated M2 (or antiinflammatory) macrophages [34]. Functionally, the M1 macrophages are mainly involved in pro-inflammatory activities, chemotaxis, and antimicrobial activities [35]. On the contrary, the M2 macrophages are engaged in the suppression of effector T cells and the promotion of tissue remodeling. Hence, they are beneficial for the resolution of inflammation and tissue repair [36]. This was also in line with our observation of macrophage subsets in the lungs of the saline group mice, where the saline group showed more and less numerous M1 and M2 macrophages, respectively, compared to those of the dexamethasone group. This strongly suggested that the polarization from M1 to M2 macrophages could play an important role in the resolution of lung injury following the administration of dexamethasone.
Surprisingly, our results revealed that, compared to the saline group, the dexamethasone group mice showed a significant increase in the granulocyte populations in the lungs. Hence, the recruitment of granulocytes, especially neutrophils, to the lungs has been considered as perhaps playing a key role in the pathogenesis of the progression of lung injury and targeting granulocyte recruitment has recently been considered as a therapeutic strategy [37,38]. At first glance, these data seem to contradict our observations. However, other studies have provided evidence that, after the occurrence of inflammatory conditions, the neutrophil phenotype can be changed from a pro-inflammatory type to an anti-inflammatory type through the production of pro-resolving mediators, such as lipoxins, after which inflammation is resolved [39]. Importantly, our findings revealed a negative correlation among the granulocyte populations but positive correlations for other studied immune cell populations (macrophages and T-and B-lymphocytes) within the lungs and MFALCs. This suggests that the latter immune cell types within the MFALCs may have a role in the pathogenesis of lung injury rather than the granulocytes. Moreover, our results revealed a significant reduction in the degree of the proliferation of immune cells following dexamethasone administration, which suggested another postulating ameliorative effect of dexamethasone on the degree of lung injury in the Lpr mice characterized by lymphoproliferative disorder [40]. Interestingly, our results also revealed significant alleviation in the expression of lung CXCL9 in the dexamethasone group compared to that of the saline group. CXCL9 plays an essential role in the induction of the chemotaxis, differentiation, and proliferation of leukocytes [21]. Therefore, we postulated that the impact of dexamethasone on resolving immune cell proliferation from an injured lung could be via the reduction in the expression of CXCL9 and its proliferation tendency.

Experimental Animals and Ethical Statement
Female Lpr mice (aged 12 weeks) were purchased from Japan SLC Inc. (Shizuoka, Japan) and maintained in specific pathogen-free housing conditions. The researchers adhered to the Ethical Guidelines for Laboratory Animal Care and Use, which was approved by the Institutional Animal Care and Use Committee of the Graduate School of Veterinary Medicine, Hokkaido University (Approval No. 15-0079).

Experimental Design and Sample Preparation
Administration of dexamethasone was performed according to our previous study [17]. Briefly, the Lpr mice were divided into the following two groups: the dexamethasone group and the saline group. The former group received dexamethasone with drinking water as a daily oral dose along with weekly intraperitoneal injections (for 10 weeks) at a dose of 0.4 mg/kg of body weight. The same protocol was followed for the latter group, but the dexamethasone was replaced with saline. At 22 weeks of age, deep anesthesia was administered to all the mice of both groups (intraperitoneal injection of 0.2 mL anesthetic solution "mixture of 5.0 mg/kg B.W. butorphanol, 4.0 mg/kg B.W. midazolam, and 0.3 mg/kg body weight medetomidine"/10 gm body weight). Next, the blood was collected from the femoral artery. Later, the head of the mouse was dislocated, and both the lung and mediastinal fat tissues (MFTs) were immediately collected and immersed in 4% paraformaldehyde for overnight fixation at 4 • C. The tissues were then processed for paraffin embedding.

Serum Autoantibody Measurement
To examine the severity of systemic autoimmune disease, the levels of serum anti-dsDNA abs were used as the index of autoimmunity, this was measured using an ELISA kit (FUJIFILM Wako Pure Chemical Corporation, Osaka, Japan) as per the manufacturer's instructions.

Tissue Preparation for Histopathological and Immunohistochemical Analysis
Paraffin sections of 3 µm were prepared from MFT and lung tissues. These sections were subjected to either hematoxylin and eosin (HE) staining for routine histopathological examination or immunostaining according to our previous study [11].
Immunohistochemical staining was performed to detect the immune cells (B220+ Blymphocytes, CD3 + T-lymphocytes, Iba1 + macrophages [41][42][43], and Gr1 + granulocytes), proliferating cells (Ki67), and the vessels (peripheral node addressin "PNAd", high endothelial venules "HEVs", and lymphatic vessel endothelial hyaluronic acid receptor 1 "LYVE-1" lymphatic vessels "LVs"). After deparaffinization and rehydration, heat-induced antigen retrieval was applied to the paraffin sections. The sections were incubated in absolute methanol (containing 3% hydrogen peroxide) for 20 min at room temperature to quench the endogenous peroxidase activity. This was followed by flushing the sections with distilled water (DW). Then, the sections were incubated with normal serum for 1 h at room temperature to block the non-specific binding sites. After blocking, the sections were incubated overnight with specific primary antibodies at 4 • C (Table 1). Next, the sections were rinsed with phosphate buffer saline (PBS) 3 times, with each rinse performed for 5 min. The washed sections were incubated with specific biotinylated secondary antibodies for 30 min at room temperature. Later, the sections were rinsed with PBS (3 times/5 min each) and incubated with streptavidin for 30 min at room temperature. The sections were again washed with PBS, and the positive reactions were developed by adding 3, 3 diaminobenzidine tetrahydrochloride-H 2 O 2 solution for a short incubation time. Next, the nuclei were counterstained with hematoxylin. Finally, the sections were dehydrated, cover slipped, and examined under a microscope.

Morphometrical Measurements
NanoZoomer 2.0-RS (Hamamatsu Photonics, Shizuoka, Japan) was used to prepare the virtual slides of HE-stained lung and MFT sections to analyze the percentages of the ratios for the area of the MNCA/area of the lung field and the area of LCs/the area of total MFTs between the study groups. The NDP view2 software (Hamamatsu Photonics Co., Ltd., Hamamatsu City, Japan) was used to measure the area of MNCA/area of the lung field and the area of the LCs/area of the total MFTs, after which the percentages of the averages were compared between the study groups. Similarly, the NDP view2 software was also used to measure the ratios of HEVs and the area of LVs/both the total lung field and the area of MFALCs in the virtual slides, converted to immuno-stained PNAd and LYVE-1 lung and MFT sections, which were then compared between both groups.
To measure the positive index ratio of the immune cells (CD3 + T-lymphocytes, B220 + B-lymphocytes, Iba1 + macrophages, and Gr1 + granulocytes), we captured digital images of immunohistochemically stained lungs and MFTs at 20× magnification using the BZ-X710 microscope. Next, we measured both the field area and immune cell positive area within the corresponding field area using a BX-analyzer (Keyence, Osaka, Japan). Then, the average ratios of immune cells positive area/field area were calculated and compared between the study groups. A BX-analyzer (Keyence) was also used to quantify the degree of proliferation, the ratio of Ki67 positive cell density (number/mm 2 ), and the number of Ki67 + cells/the total number of immune cells in the lungs and MFALCs. Similarly, the cell densities of CXCR5 + and CXCL13 + cells in the lung sections were measured, and the averages were compared between the study groups.

Statistical Analysis
The statistical analyses among the study groups were conducted using the Graph pad Prism 6 software (GraphPad Software, San Diego, CA, USA). The differences among different numerical results in the saline and dexamethasone groups were compared using the Mann-Whitney U test. The data were presented as mean ± standard error (SE). Significant (*) and highly significant (**) differences were considered at p values < 0.05 and <0.01, respectively. Additionally, the correlation coefficients between two different parameters were determined using Spearman's correlation test (p < 0.05).


s2fieldsofstudy: Biology Chemistry

text:
Modular Site-Specific Conjugation of Nanobodies Using Two Co-Associating Tags

The homogeneous labeling of antibodies and their fragments is a critical step for the generation of robust probes used in immuno-detection applications. To date, numerous chemical, genetic and peptide-based site-specific coupling methods have been developed. Among these methods, co-assembling peptide-tags is one of the most straightforward and versatile solutions. Here, we describe site-specific labeling of nanobodies through the use of two co-associating peptides tags, E3 and K3, originating from the tetramerization domain of p53. These E3 and K3-tags provide a simple and robust method for associating stoichiometric amount of VHH and fluorescent probes, either fluorescent proteins or fluorochromes, at specific positions. As a proof of concept, a nanobody targeting the human epidermal growth factor receptor 2 (HER2), the nano-HER2 was genetically fused to the E3 and associated with different fluorescent K3-derivates. Entities were produced separately in Escherichia coli in soluble forms at high yields and co-assembled in vitro. These molecular probes present high binding specificity on HER2-overexpressing cells in flow-cytometry with relative binding constants in the low nanomolar range and are stable enough to stain HER2-receptor on living cells followed detection using fluorescent confocal microscopy. Altogether, our results demonstrate that the non-covalent conjugation method using these two co-associating peptides can be easily implemented for the modular engineering of molecular probes for cell immuno-staining.

Introduction
Antibodies and their derived fragments are essential tools to visualize cellular proteins and cell surface receptors. To this regard, labeling of these binders is a critical step for the generation of high quality and homogenous probes suitable for various applications such as fluorescence-microscopy, flow cytometry, diagnostic, therapeutic applications and molecular imaging. A very promising alternative to antibodies, Fabs and scFvs is the use of variable domains of heavy chain antibodies from Camelidae (V H Hs or nanobodies). Indeed, V H Hs are highly soluble, stable under various physicochemical conditions and can be produced at high yield in E. coli. They can also be easily engineered, opening a wide range of labeling strategies [1][2][3][4].
V H Hs can be labeled either covalently or non-covalently with fluorochromes or fluorescent proteins [5]. However, each of these methods suffers from several drawbacks. For instance, chemical methods involving the covalent conjugation to either side chain ε-amine group of a lysine or sulfhydryl group of a cysteine may alter residues inside or near the antigen-binding site, leading to the generation of non-functional reagents. Maleimide conjugation to cysteine residues can lead to labeling heterogeneity and in some cases to irreversible unfolding, especially if buried scaffold cysteine residues are conjugated, resulting in a heterogeneous population of labeled tracer [5]. Therefore, several alternatives have been developed to circumvent these shortcomings. Thus, introduction of an additional unpaired cysteine at the carboxyl-terminal end of the nanobody for site-specific conjugation via a thio-ether bond has proven to be a reliable method for the generation of homogenous tracers [6]. This approach has been further developed on an anti-Nup98 nanobody in which cysteine residues were engineered at the nanobody surface for site-specific conjugation, leaving the internal framework cysteine residue fully intact [5]. But to better guide cysteine assignment in the nanobody framework, the crystal structure of the anti-Nup98 nanobody in complex with the Nup98 protein had to be solved, making this approach cumbersome for its application on different nanobodies.
Apart from the genetic engineering of unpaired cysteine residue for site-specific labeling, other alternatives explored peptide-tag approaches for site-specific protein labeling. Enzyme-based tags provide high specificity and enable fast covalent site-specific labeling reactions [7]. Peptide-based recognition tags are categorized into three groups. First, the chemo-enzymatic labeling techniques, such as biotin ligase, transglutaminase or Sortase A, can achieve site-specific labeling of a protein of interest in one or two coupling steps [7]. For example, a short tubulin-derived recognition sequence (Tub-tag) was used successfully for C-terminal labeling of nanobodies [8]. Moreover, the protein-ligation capacity of the enzyme Sortase A allowed site-specific coupling of three different probes to an anti-HER2 nanobody for in vivo imaging applications [3]. However, to achieve high labeling yields, long reaction times and high concentrations in the µM range of substrates were required. For the Sortase A reaction, nucleophilic probes have to be synthetized, and the overall efficiency of the conjugation procedure does not exceed 30 to 50% [3]. The second group of peptide-based tags is set on metal ion-dependent labeling with cysteine-rich tags or metal-chelating tags such as His-tags, allowing one-step labeling. However, cytotoxicity, background signals due to non-specific interactions with other proteins and non-specific binding are the main drawbacks [7]. The last group consists of peptide-binding peptide tags. One example is the artificial heterodimeric coiled-coil peptides [9]. These tags, which form heterodimers with high affinity in the nM range, were, however, not tested for site-specific labeling of V H Hs [10].
Beside chemical labeling and peptide tags, a vast number of fluorescent proteins (FPs) with a broad color palette spanning nearly all visible and far-red spectral regions are available [11]. Nanobodies genetically fused to these fluorescent encoded probes are called chromobodies and have been successfully expressed in living cells to trace intracellular proteins using fluorescence microscopy, such as Dmnt1 or microtubules [12,13]. To date, one anti-CD38 chromobody genetically fused either to eGFP or to mCherry has been used successfully for flow cytometry [14].
In this report, we describe a straightforward modular strategy for a site-specific conjugation of nanobodies suitable to either fluorochromes or FPs using the co-associating peptides E3 and K3 [15][16][17][18]. These two short peptidic sequences of 31 amino acid residues, which originate from the tetramerization domain of p53 (residues 325-355), co-associate exclusively through helix-helix contacts to form a dimer of dimers. The α-helical charged interface involving lysine and acid glutamic residues modulates tetramer stability through salt bridges. We describe the use of E3 and K3 peptides to engineer bifunctional site-specific labeled macromolecules for immuno-detection of receptor proteins in fluorescence microscopy and in flow cytometry. As a proof of concept, the HER2 receptor was targeted by a nanobody anti-HER2 fused to the E3 peptide (hereinafter referred to as nano-HER2-E3) coassembled with either the labeled synthetic K3 peptide or with the K3 peptide genetically fused to various FPs and expressed as recombinant proteins. Our data clearly demonstrate that this non-covalent labeling strategy allows the formation of robust and modular site-specific labeled probes for immuno-detection of receptor proteins in fluorescence microscopy or flow cytometry.

Results
The bivalent nano-HER2-E3, already described in our previous studies [16], was evaluated as a molecular probe for imaging and flow cytometry. Conjugation of the nano-HER2 with Alexa-Fluor-488 dye via lysine residues using N-hydroxysuccinimide (NHS) esters has been shown to result in a strong affinity drop of 20 to 1000 fold [16,19]. Thus, as the nanobody contains two cysteine residues and its oxidative status appeared not to be critical for epitope recognition [20], we tried to perform non-specific labeling on cysteine residues using maleimide conjugation. However, the degree of labeling was extremely low and could not be implemented for imaging and flow cytometry.
A cysteine residue was then introduced at the C-terminal part of the nano-HER2-E3 and used as unpaired cysteine for covalent maleimide conjugation using an Alexa-Fluor-488 dye (referred to as nano-HER2-E3-C). The nano-HER2-E3-C was then produced in E. coli, but at lowest yields compared to the parental counterpart, as previously observed with other nanobodies [2]. Introduction of a cysteine residue caused extensive dimerization as shown by the size-exclusion chromatography ( Figure S1A,B), where the nano-HER2-E3-C eluted into two peaks without a reducing agent, TCEP. Thus, reduction using TCEP was needed prior labeling with maleimide conjugate Alexa-Fluor-488 dye, but the degree of labeling was modest and never reached more than 70%, leading to a heterogeneous labeling population.
Therefore, we decided to evaluate the co-associating tags as a labeling device. Nano-HER2-E3 was evaluated for co-assembly with various fluorescently labeled K3 peptides. To that end, the K3 peptide was either genetically fused to a C-terminal part of FPs such as eGFP or mScarlet (hereafter referred to as eGFP-K3 and mScarlet-K3, respectively) or chemically synthetized with an additional cysteine residue (hereinafter referred to as C-K3) ( Figure 1). We took advantage of the absence of cysteine residue in the K3 peptidic sequence to introduce a N-terminal cysteine residue that can be selectively conjugated to maleimide-functionalized fluorochromes. The different recombinant proteins were produced at extremely high levels in E. coli and subsequently purified by immobilized metal affinity chromatography followed by sizeexclusion chromatography. The final yields of all purified proteins were above 100 mg/L. Elution profiles revealed that all K3-fused displayed profiles corresponding to tetramers with a size between 155 and 160-kDa (Table S1), whereas nano-HER2-E3 behaved as dimers, as previously described [16]. In addition, the elution profile of the different proteins on size-exclusion chromatography showed a major single peak, indicating an oligomerization of almost 100%.
We next evaluated the ability and the stoichiometry of the complex formation between nano-HER2-E3 and its various cognate K3 partners. Following western blot transfer, the K3 moieties were detected by direct fluorescence using a Typhoon TM device, whereas the nano-HER2-E3 was detected using an anti-c-myc monoclonal antibody followed by a secondary anti-mouse mAb either IR700-labeled (red) or IR800-labeled (green). Increasing amounts of purified nano-HER2-E3 were mixed with steady amounts of eGFP-K3 or Alexa-Fluor-647-labeled synthetic K3 peptide (hereinafter refereed as to 647-C-K3). Mixtures were analyzed by native polyacrylamide gel electrophoresis ( Figure 2). The free eGFP-K3 (green signal) was gradually titrated by increasing amounts of nano-HER2-E3 (red signal) leading to the formation of yellow fluorescent bands corresponding to the heterotetramers (Figure 2A). At a molar ratio of 1:1, the eGFP-K3 was entirely complexed to the nano-HER2-E3. By comparison, the same experiment conducted with eGFP-E3 instead of eGFP-K3 showed no association with nano-HER2-E3 ( Figure 2B). As expected, the free homodimer eGFP-E3 migrated faster in the native gel electrophoresis than the homotetramer eGFP-K3. Likewise, the 647-C-K3 synthetic peptide (red signal) displayed a strong size shift in native polyacrylamide gel electrophoresis subsequent to its co-association with the nano-HER2-E3 (green signal) ( Figure 2C). Furthermore, eGFP-K3 was entirely titrated when mixed with an excess of mCherry-E3 in 85% fetal bovine serum (FBS), confirming a strong and highly specific interaction between E3 and K3 peptides independently of the fusion partner ( Figure S2). Thus, the visualization of these complexes clearly demonstrates that the E3 peptide efficiently co-associates with its K3 cognate peptide partner fused to the eGFP or labeled with Alexa-Fluor-647 dye.
We confirmed the efficiency of the heterotetramerization by analytical size-exclusion chromatography (SEC) ( Figure S3). When equimolar amounts of nano-HER2-E3 and eGFP-K3 were mixed and analyzed by SEC, new complexes were observed in both cases. These new complexes showed intermediary molecular weights compatible with the theoretical molecular weight of the heterotetramer ( Figure S3A). In addition, elution profiles established that the homotetramer eGFP-K3 dissociates to the benefit of more stable heterotetrameric complexes formed with nano-HER2-E3. For the heterotetramer nano-HER2-E3/647-C-K3, the SEC was not resolutive enough and only a very slight difference in molecular weight could be observed ( Figure S3B). However, a SEC could clearly separate nano-HER2-E3/647-C-K3 from the free peptide 647-C-K3. Overall, the formation of the complexes was up to 95%, as no residual homo-oligomers could be detected.
After the initial in vitro validation, we decided to evaluate the close interaction between E3 and K3 moieties at the cell surface by using Försters resonance energy transfer (FRET) ( Figure 3). As energy transfer in FRET requires proximity of less than 10 nm between the donor and the acceptor, this assay can confirm the association between the nano-HER2-E3 and the various fluorescent K3 derivatives. Two fluorophores suitable for FRET were chosen to perform flow cytometry, with Alexa-Fluor-488 as the donor and mScarlet as the acceptor ( Figure 3A). The nano-HER2-E3-C-488 co-associated with mScarlet-K3 was tested by flow-cytometry on two breast cancer cells lines, HER2-positive HCC1954, HER2-negative MDA-MB-231 [21] and on HER2-silenced HCC1954 cells using small-interfering oligonucleotides ( Figure 3A,B and Figure S4A,B). The fluorescent moieties were excited at 488 nm, and the signal was recorded through their respective filters, 533/30 for Alexa-Fluor-488 and 585/40 for mScarlet. As expected, a clear signal was detected around 525 nm on HER2+ HCC1954 cells incubated with nano-HER2-E3-C-488 with a mean fluorescence intensity (MFI) of 5258 ( Figure 3B), whereas no detectable signal was observed on HER2-MDA-MB-231 or on HER2-silenced HCC1954 cells under the same conditions ( Figure S4A,B). For the nano-HER2-E3/mScarlet-K3 complex, only a weak signal was observed on HCC1954 cells (MFI of 351), as the mScarlet is weakly excited at 488 nm ( Figure 3A,B). However, when the two fluorophores were paired in the nano-HER2-E3-C-488/mScarlet-K3 complex, a clear increase of the mScarlet signal intensity could be observed on HCC1954 cells. The mScarlet MFI was 982 and was accompanied with a decrease in the Alexa-Fluor-488 signal intensity (MFI of 3431), as expected following energy transfer from the donor to the acceptor. These observations indicate that FRET is occurring within the nano-HER2-E3-C-488/mScarlet-K3 complex, confirming the efficient co-association between nano-HER2-E3-C-488 and mScarlet-K3 at the cell surface.  In order to validate the results of our flow cytometry-based FRET assay in an independent setting, we performed confocal microscopy. Heterotetrameric complexes with distinct fluorophore pairs, nano-HER2-E3-C-488 co-assembled with mScarlet-K3, were incubated either directly in the culture media of the HCC1954 cells for 30 min at 37 • C or after fixation of the cells. In both cases, 100% of the cells showed an intense and distinguishable staining at the cell membrane and displayed a high amount of color-coded FRET ( Figure 3C), confirming the results obtained with our FACS-based FRET assay ( Figure 3A,B) and clearly demonstrating the close proximity of E3 and K3 partners. In addition, the strong FRET signal on living cells experiment proves the stability of the E3-K3 interaction even in culture media. No signal could be detected on MDA-MD-231 control cells, nor with nano-HER2-E3-C-488 or nano-HER2-E3/mScarlet-K3 alone, confirming the specificity of the signal ( Figure S5A). Similar results were obtained with the fluorophore pairs, nano-HER2-E3-C-568 co-assembled with eGFP-K3 ( Figure S5B). After this validation, we wanted to determine if our labeling strategy could be comparable and therefore potentially substitute the classical indirect immunofluorescence protocol. Nano-HER2-E3 co-associated with either 488-C-K3 or eGFP-K3 was compared to an anti-HER2 mouse mAb, followed by a secondary Alexa-Fluor-488 mAb. No obvious differences could be observed between both methods ( Figure S6).
Finally, we wanted to assess whether the formation of these complexes via E3 and K3 peptides affected the binding efficiency of the nano-HER2 at the cell surface HER2 receptor. To that end, the relative binding constants of the nano-HER2-E3, either alone or co-associated with the synthetic peptide 488-C-K3 or the eGFP-K3, were assessed on HER2 + HCC1954 and HER2 -MDA-MB-231 breast cancer cells by flow cytometry (Figure 4). As shown in Figure 4B, association of either eGFP-K3 or K3-C-488 with nano-HER2-E3 led to only a five-fold decrease in the relative binding affinity compared to nano-HER2-E3-C-488. Indeed, the relative binding efficiency was 6.3 ± 1.8 nM for nano-HER2-E3-C-488, as previously estimated [16]; 24.3 ± 7.0 nM for the complex nano-HER2-E3/eGFP-K3; and 28.7 ± 8.1 nM for the nano-HER2-E3/K3-C-488 heterotetramer. No signals were observed on HCC1954 cells incubated with nano-HER2/eGFP-K3, confirming the strict dependency between the fluorescent positive signal and the co-association of nano-HER2 with fluorophore via the E3 and K3 peptides ( Figure 4B). Moreover, no signals were detected on HCC1954 (HER2 + ) cells incubated with a mock nanobody directed against the GFP protein (nano-eGFP-E3), described elsewhere [18], or on MDA-MB-231 (HER2 -) cells, demonstrating the binding efficiency and specificity of the nano-HER2-E3 and fluorescent K3 complex for the HER2 cell surface receptor.

Discussion
Numerous labeling methods have been developed to conjugate fluorochromes or fluorescent proteins to antibody fragments, such as scFv or V H Hs [13,22,23]. Conventional bioconjugation of an organic fluorophore to lysine or cysteine residues of an antibody or derivatives suffers several drawbacks. First, an indiscriminate coupling reaction with respect to the target amino acid residue can lead to partial or even complete disruption the antigen-binding site, resulting in the generation of non-functional probes. In addition, chemical conjugations generally result in a heterogeneous mixture of antibody fragments having different number of fluorochromes per molecule. Thus, defining the coupling site is a key goal to overcome these limitations [6].
Here, we described a novel peptide tag-based labeling method taking advantage of a strong, non-covalent, highly specific conjugation between two short co-associating peptides, E3 and K3, for site specific labeling. In our approach, the E3 tag was introduced at the C-terminal end of the anti-HER2 V H H, giving rise to nano-HER2-E3, and the K3 peptide was either chemically synthetized with a modifiable amino acid or genetically fused to the N-terminal of various FPs. By separating the molecular probe into two distinct entities, our system is able to overcome yield, folding and solubility issues, which can be major problems when producing chimera-encompassing FPs in bacteria. For instance, the chromobody V H H-CD38-eGFP expressed in E.coli was mostly soluble, but with a yield of only around 6 mg of pure recombinant protein per liter of culture [14]. In our case, the strategy to separate both entities is illustrated by the excellent yields of the production: up to 100 mg per liter for the V H H and more than 120 mg per liter of culture for both eGFP-K3 and mScarlet-K3.
Then, association of both partners takes place in few minutes at RT or on ice, in a high concentration of serum and at nanomolar concentration. Unlike metal-ion peptide tags, no apparent toxicity and unspecific binding were noticed with our system. This technique is particularly straightforward and convenient, and co-assembly occurs efficiently with different fluorescent partners, such as Alexa-Fluor maleimide K3 peptide or FPs genetically fused to K3 peptide.
In addition, this technology allows a great flexibility in the choice of the fluorescent partner to be associated. Several fluorescent K3 probes were tested in association with nano-HER2-E3 without loss of binding specificity and efficiency. An indirect advantage of the system is that bivalency of the nano-HER2 is also achieved by the dimerization of the E3 tag, improving its functional affinity compared to the monovalent counterparts, by mean of avidity [18].
Analysis of the association between the V H H fused to E3 and the K3 derivate-labeled peptides demonstrate that pairing preferences led exclusively to the rapid formation of heterotetrameric complexes to the disadvantage of the homodimer and homotetramer monovalent counterparts. Complexes were remarkably stable during gel electrophoresis and in size-exclusion chromatography. Using FACS and confocal fluorescent microscopy, we showed that our bio-conjugated probes are highly stable in culture media to stain specifically the membrane of HER2 positive cells. In confocal experiments, the signal given by the various probes incubated before fixation of the cells was comparable to the signal detected after incubation of the probes on fixed cells. These results were further corroborated with the strong FRET signal, demonstrating that at least at the cell membrane E3 and K3 are tightly associated. Another advantage of these molecular probes is to avoid the use of secondary labeled antibodies and extensive washing steps, simplifying the procedure and reducing potential background signal. Finally, the nano-HER2 heterotetramers demonstrated equivalent apparent binding efficiency on HCC1954 cells as the nano-HER2 homodimer, demonstrating that co-association of both partners does not affect the folding and/or the binding efficiency.
In conclusion, we have developed a highly efficient and specific tool for site-specific labeling of nanobodies resulting in a homogeneous tracer population. Our system can be seen as a "two-in-one" modular site-specific labeling system. It takes advantage of the dimerization E3 tag to increase the affinity of the nanobody by means of avidity, and uses it as an anchor for its co-associating partner, K3, for a labeling device. Moreover, connecting E3 and K3 peptides to macromolecules does not compromise the activity of the fusion partners and can be easily achieved by genetic engineering or chemically. This straightforward and inexpensive technology can be scaled up for mass production of nanobody-based reagents. Finally, tethering two distinct modular entities into molecular probes allows a great flexibility in terms of co-associating partners. Altogether, E3 and K3 tags represent a versatile methodology for conjugating macromolecular probes or different chemical moieties to a protein of interest.

Plasmids
Nano-HER2-E3 expression plasmid has been described elsewhere [16,18]. The K3 coding sequence was introduced into the NheI-EcoRI restriction sites of pET-eGFP-E3 to replace the E3 sequence, leading to pET-eGFP-K3 [15]. A sequence encoding for mScarlet and optimized for E. coli expression was synthetized by Integrated DNA Technologies (IDT), amplified by PCR and cloned into NcoI-NheI restriction sites in frame with K3 sequence of the pET-eGFP-K3.

Expression and Purification of the Recombinant Fusion Proteins
Briefly, E3 and K3 fusion proteins were overexpressed in E. coli BL21 (DE3) pLysS after induction with 0.5 mM isopropyl thiogalactoside (IPTG). After 24 h at 20 • C, the cells were harvested, re-suspended in 20 mM Phosphate buffer pH 8, 250 mM NaCl and 10 mM imidazole. Following lysis with lysozyme and sonication, cell debris was removed by ultra-centrifugation at 36,000× g, and the supernatant was applied to IMAC chromatography column charged with cobalt (GE Healthcare Saclay, Paris, France). IMAC purified fractions were subsequently loaded on a HiLoad 16/60 Superdex 200 prep grade column (GE Healthcare, Bio-sciences AB, Uppsala, Sweden) operating at a flow rate of 0.5 mL/ min. Aliquots of the fractions were separated on SDS-PAGE gels and analyzed by Coomassie blue staining. Complex formation between labeled recombinant proteins was controlled by analytical chromatography on Superdex 200 Increased 5/150 GL (GE Healthcare, Bio-sciences AB, Sweden).

Transient siRNA Transfections
Transient siRNA transfections were performed using Lipofectamine RNAiMAX (Invitrogen, P/N 56532, Carlsbad, CA, USA) according to the manufacturer's instructions. HER2-targeting siRNAs were SMARTpoolON-TARGETplus obtained from Dharmacon. For controls, siRNAs ON-TARGET plus non-targeting pool from Dharmacon were used. siRNAs were used at a final concentration of 10 nM, and cells were transfected 24-72 h prior to experiments.

Fluorescence-Activated Cell Sorting (FACS)
Trypsinized cells were incubated with a mixture of equimolar ratio of recombinant proteins as indicated, for 30 min at 4 • C in FACS Buffer (PBS with 0.5% BSA and 2 mM EDTA). Cells were washed twice in FACS Buffer and analyzed on an Accuri™ C6 Plus flow cytometer (BD Bioscience, San Jose, CA, USA). The relative mean fluorescence intensities were normalized and plotted against the concentration of the nano-HER2 at monomer concentration. The data shown here are single-point measurements.

Immunofluorescence
The cells grown on coverslips were either incubated with the labeled complex and fixed with 4% (w/v) paraformaldehyde for 30 min or directly fixed and permeabilized with 0.2% Triton X-100 for 5 min before incubation with the labeled complex or with a commercial anti-HER2 mouse mAb (9G6) was used for indirect immunofluorescence (Thermofischer, Carlsbad CA, USA). The coverslips were mounted with Fluoromount G containing 4',6'-diamidino-2 phenyleindole (SouthernBiotech, Birmingham, UK). Images were captured with deconvolution (confocal) fluorescence microscopy (Leica DMIRBE inverted microscope and OPENLAB 3.1.4 software, Nussloch, Germany) and images were processed using Fiji. Images were filtered by applying the difference between two Gaussian Blurs (1.5 and 25).

Native-PAGE
Separation of proteins by non denaturing polyacrylamide gel was performed, as previously described [15]. Shortly after this, purified complexes were mixed as indicated, and resolved on nondenaturing polyacrylamide gradient gel (8−18%) at pH 8.8. Proteins were transferred on nitrocellulose membrane and visualized using a direct fluorescence Typhoon FLA 9500 biomolecular imager (GE Healthcare) at 555 nm or detected with an anti-myc mAb and a secondary antibody IRDye800-conjugated anti-mouse (Science-Tec) and visualized on an Odyssey Classic Infrared Imager (LI-COR).


s2fieldsofstudy: Biology Medicine

text:
Protective Roles of Apigenin Against Cardiometabolic Diseases: A Systematic Review

Apigenin is a flavonoid with antioxidant, anti-inflammatory, and anti-apoptotic activity. In this study, the potential effects of apigenin on cardiometabolic diseases were investigated in vivo and in vitro. Potential signaling networks in different cell types induced by apigenin were identified, suggesting that the molecular mechanisms of apigenin in cardiometabolic diseases vary with cell types. Additionally, the mechanisms of apigenin-induced biological response in different cardiometabolic diseases were analyzed, including obesity, diabetes, hypertension and cardiovascular diseases. This review provides novel insights into the potential role of apigenin in cardiometabolic diseases.

INTRODUCTION
Apigenin (4 ,5,7-trihydroxyflavone) is named after the genus Apium belonging to family Apiaceae (1). It is widely distributed in vegetables and fruits, such as celery, parsley, oranges and garlic (2), and is also found in herbs such as snow lotus and chamomile (3) (Figure 1A). As a secondary plant metabolite, apigenin is usually stored in plants in a water-soluble glycosylated form (4). Purified apigenin is a yellow powder with a low molecular weight (MW 270.24). It is nearly insoluble in water, moderately soluble in hot alcohol and soluble in dimethyl sulfoxide (DMSO) (5). Pure apigenin is chemically unstable and therefore stored in the dark at −20 • C (5).
The biosynthesis of apigenin occurs on the surface of the endoplasmic reticulum and requires four steps including intermediate synthesis, basic skeleton synthesis, precursor synthesis and generation of the apigenin structure (6) (Figure 1B). Current evidence indicates that the bioactivity of apigenin is dependent on its chemical structure, thus the structure-activity relationship of apigenin can be determined by extracting the molecular fragments associated with a specific biological activity. For example, double bonds in the two aromatic rings and hydroxyl groups on C-7 and C-4 induce the inhibition of α-glucosidase and α-amylase (7). The C-4 hydroxyl group in ring B is essential for immunomodulatory properties (8). The hydroxyl radicals at position 5, 7 and 4 are necessary for Liver X receptor activation (9).
Cardiometabolic disease links the metabolic syndrome disorders (abdominal adiposity, hypertension, dyslipidemia, hyperinsulinemia and glucose intolerance) that are predictive of cardiovascular disease and Type 2 diabetes (T2DM) (10). Recently, apigenin has been found to play a protective role in cardiometabolic diseases in vitro and in vivo. This systematic review summarizes the current perspective.

PROTECTIVE ROLES OF APIGENIN IN CARDIOMETABOLIC DISEASES Protective Role of Apigenin in Obesity and Lipid Metabolism
Obesity is attributed to chronic energy imbalance, including excessive energy intake and limited energy expenditure (11,12). Anti-obesity strategies focus on suppression of energy intake and stimulation of energy expenditure by regulating lipid metabolism, such as inhibiting pancreatic lipase activity and adipocyte differentiation (13). Studies suggest that apigenin controls energy intake by inhibiting appetite and stimulating energy expenditure by regulating lipid metabolism to alleviate obesity.
First, apigenin inhibits obesity by suppressing food consumption. In vitro and in vivo studies confirmed that apigenin upregulates the expression of anorexigenic neuropeptides pro-opiomelanocortin (POMC) and cocaine-and amphetaminerelated transcript (CART), resulting in inhibition of food intake. N-29-2 and SH-SY5Y cells transfected with pPOMC-Luc and pCART-Luc vectors were treated with 0.2-5 µM apigenin for 6 h, resulting in upregulation of pPOMC-Luc and pCART-Luc activity. In an in vivo study, 6-week-old C57BL/6J mice were injected (every 24 h) intraperitoneally with 1 or 10 mg/kg of apigenin in a short-term intervention. Male C57BL/6J mice fed with a high-fat diet (HFD) or a standard laboratory chow diet received 0.05% apigenin for 30 days to demonstrate that apigenin reduces food intake and visceral fat over a long-term period (14). POMC and CART neurons found in the retro-chiasmatic area and throughout the rostrocaudal span of the arcuate nucleus (ARC) play a role in appetite control (15). Increasing expression of POMC and CART induces the expression of leptin receptor B (LepRb) to facilitate leptin binding to LepRb, resulting in an anorexic effect and upregulation of insulin receptors to inhibit appetite.
Secondly, apigenin stimulates energy expenditure by regulating lipid metabolism, including adipogenesis, lipolysis, fatty acid oxidation, and cholesterol synthesis. Recent studies indicate that adipose tissues are generally targeted by apigenin eliciting the following effects: (1) Stimulation of PPARγ signaling. Several studies have demonstrated that apigenin inhibits adipocyte differentiation via STAT3 (the signal transducer and activator of the transcription 3)-CD36-PPARγ (peroxisome proliferator-activated receptorgamma) axis (16) and AMPK (5 -Adenosine monophosphateactivated protein kinase)/PPARγ axis (17). One study showed that 100 µM apigenin treatment inhibits the differentiation of 3T3-L1 preadipocytes to mature white adipocytes. Mouse models of diet-induced obesity receiving apigenin via subcutaneous injection for 13 days showed that apigenin reduced visceral fat mass (16). Apigenin binds to non-phosphorylated STAT3 to decrease STAT3 phosphorylation and nuclear translocation (18), followed by a decline in the expression of CD36, the downstream target gene involved in fatty acid transport (19). PPARγ is the transcript factor of central ligand-activated transcription factors. It inhibits adipogenesis and controls adipose tissue differentiation to regulate inflammation in obesity. PPARγ expression depends on CD36 expression and therefore apigenin treatment inhibits adipocyte differentiation via downregulation of PPARγ. Other studies reported that apigenin activates the phosphorylation of AMPK (5 -adenosine monophosphate-activated protein kinase) to downregulate adipogenesis via AMPK/PPARγ axis in 3T3-L1 cells treated with 10 µM apigenin for 1 h or 4 days (17) and in HFD mice treated with 200 mg/kg enzyme-treated celery extract (20). AMPK acts as a potential target against adipogenesis (21,22) and downregulates the expression of PPARγ. The adipogenous genes downstream of PPARγ, such as fatty acid-binding protein 4 and stearoyl-CoA desaturase, are also downregulated, thereby suppressing adipogenesis (17).
(2) Repression of enzyme activity. Guo et al. reported that 0.6 mM apigenin directly inhibits pancreatic lipase activity in vitro (23). Pancreatic lipase catalyzes the conversion of triglycerides to monoglycerides and fatty acids in the intestine. Obesity is alleviated by the suppression of pancreatic lipase, fatty acid synthesis and fat absorption. Gómez-Zorita et al. showed that treatment with 25 µM apigenin decreases the expression of fatty acid synthase (FAS), while increasing the expression of adipose triglyceride lipase (ATGL) in mature adipocytes derived from human mesenchymal stem cells (hMSCs), resulting in reduced adipogenesis (24).
Liver, in addition to adipose tissues, is essential for lipid metabolism. Abnormal lipid metabolism in the liver induced by obesity may cause hepatic steatosis. Apigenin also improves lipid metabolism in the liver to alleviate hepatic steatosis via following mechanisms: (1) Stimulation of PPARγ signaling. Apigenin modulates PPARγ expression in hepatic lipid metabolism via Nrf2-PPARγ axis. In Hep1-6 cells, apigenin activates nuclear factor erythroid 2-related factor 2 (Nrf2) via translocation into the nucleus to upregulate downstream antioxidant enzymes and downregulate lipid synthesis (26). Activation of Nrf2 by apigenin neutralizes the activation of PPARγ to regulate lipid metabolism in liver (27).
(2) Regulation of SREBP family. Apigenin treatment inhibits lipid homeostasis by the sterol regulatory element-binding protein (SREBP) family. Apigenin significantly decrease lipid accumulation, total intracellular cholesterol (TC), and intracellular triglyceride (TG) levels via the AMPK-SREBP-1/2 (sterol regulatory element-binding protein-1/2) axis in HepG2 cells. Apigenin-induced activation of AMPK downregulates the levels of SREBP-1 and SREBP-2 to reduce the synthesis of cholesterol, fatty acids, and triglycerides in the liver. The inhibition of 3-hydroxy-3-methylglutaryl CoA reductase (HMGCR), which is the downstream target gene of SREBP-1 and FAS, the downstream target gene of SREBP-2 also regulates fatty acid and cholesterol synthesis (28).
Obesity-induced oxidative stress and inflammation also aggravate the symptoms of cardiometabolic diseases, leading to multiple cellular disorders (31,32). Current studies indicate that apigenin alleviates oxidative stress and inflammation by binding to PPARγ as an agonist to regulate M2 polarization with nuclear factor kappa-light-chain-enhancer of activated B cells (NF-κB) inhibition. Apigenin-induced PPARγ activation blocks p65 nuclear translocation. NF-κB activation is inhibited in adipose tissue macrophages, leading to an increase in M2 macrophage polarization. The anti-inflammatory effects of M2 macrophages alleviate the metabolic disorder caused by obesity-related inflammation. Meanwhile, cytokines such as IL-6, IL-1β, and TNF-α are suppressed by the inhibition of NF-κB signaling (33) Frontiers in Nutrition | www.frontiersin.org following apigenin treatment. The protective effect of apigenin on adipocyte browning in the inflammatory environment is also mediated via p65/NF-κB pathway. The inflammatory environment suppresses adipocyte browning to reduce lipid metabolism (34). Apigenin suppresses p65 translocation into the nucleus to inhibit NF-κB activation and inflammatory markers in adipocytes to attenuate inflammation and suppress adipocyte browning (25,35). Apigenin also plays a protective role in the inhibition of white-to-brown adipose tissue differentiation. The inflammation induced via activation of uncoupling protein 1 and PGE2 receptor 4 (EP4) activates the cyclooxygenase 2 (COX2)/prostaglandin E2 (PGE2) axis, resulting in conversion of white to brown adipose tissue to generate heat by excessive energy expenditure (36).
Studies reported that obesity leads to many health complications. First, obesity has been associated with gastrointestinal disorders, such as gastroesophageal reflux disease, irritable bowel syndrome, and dyspepsia (37,38). Colon inflammation adversely affects enteric motor function, leading to gastrointestinal disorders (39). Apigenin reduces the levels of malondialdehyde (MDA), interleukin-6 (IL-6), and interleukin-1β (IL-1β), as well as eosinophil infiltration in colon tissue to alleviate inflammation. Further, apigenin regulates inducible nitric oxide synthase (iNOS) expression and substance P (SP) levels in high-fat-diet (HFD)-fed obese mice (40). SP is a neurotransmitter that stimulates the contraction of various intestinal tissues and is a neurokinin receptor 1 (41). Obesity induces the expression of SP leading to enhanced tachykinergic transmission in the enteric nervous system, resulting in abnormal colonic motor function. The suppression of SP by apigenin attenuates enteric motor dysfunctions (40). NO produced by iNOS may trigger inflammation and play a role in enteric nitrergic pathways (42). The downregulation of iNOS by apigenin attenuates inflammation and enteric motor dysfunction. The regulation of gut bacteria by apigenin also prevents colonic dysfunction in mice via modulation of NOD-like receptor family pyrin domain containing 6 (Nlrp6) (43). Nlrp6 is highly expressed in the intestine. Nlrp6 deficiency may lead to proliferation of Prevotellaceae, the gut bacteria found in patients with bowel diseases (44), by promoting Nlrp6 inflammasome, IL-18 secretion, and regulation of gut bacterial homeostasis. Further, apigenin improves intestinal dysbiosis via augmentation of Akkermansia and Incertae Sedis along with reduction of Faecalibaculum and Dubosiella at the genus level (45). Obesity has also been associated with sarcopenia (46). Obesity-induced muscle atrophy also contributes to impaired glucose and lipid homeostasis, proinflammatory responses, and inflammation-induced mitochondrial dysfunction (47). Apigenin ameliorates skeletal muscle atrophy by enhancing mitochondrial function in an obese mouse model exposed to HFD and in C2C12 cells. Apigenin treatment upregulated mitochondria-related genes, including peroxisome proliferatoractivated receptor-γ coactivator-1α (PGC1α), mt-TFAM (transcription factor of PGC1α), cytochrome C, and somatic cytochrome C (CyCs) following the activation of AMPK. Such upregulation is essential for initiation of mitochondrial biogenesis and improved mitochondrial function alleviate obesity-induced skeletal muscle atrophy (48,49).
In summary, apigenin alleviates obesity and its complications via a variety of mechanisms including inhibition of appetite, glucose signaling pathways and lipid metabolism. It also regulates the intestinal microbiome, enhances mitochondrial function and attenuates inflammation and oxidative stress. The aforementioned experimental approaches and mechanisms underlying the effects of apigenin on obesity are listed in Table 1. Dosages and duration of apigenin treatment in vivo and in vitro are also listed in Table 1.

Protective Role of Apigenin in Diabetes
Diabetes also plays an important role in cardiometabolic disease (50)(51)(52). Several studies investigating the effects of apigenin on type 2 diabetes mellitus (T2DM) report decreased insulin resistance, reduced abnormal glycolipid metabolism, and alleviation of oxidative stress (53,54).
Insulin resistance plays a significant role in the pathophysiology of T2DM (55). Insulin resistance adversely affects glycometabolism in insulin-targeted organs and tissues (54). Abnormal glycolipid metabolism is a typical clinical manifestation in patients with T2DM (56). Apigenin alleviates insulin resistance and glycolipid metabolic disorders via following mechanisms: (1) Inhibition of insulin receptor kinase. Apigenin inhibits tyrosine nitration of the insulin receptor kinase domain leading to alleviation of insulin resistance. Tyrosine nitration of IRβ (intracellular β subunits of the insulin receptor) may lead to decreased tyrosine phosphorylation, resulting in impaired insulin signal transduction in HFD mice (57). In vitro studies showed that apigenin decreases the Cu 2+ -catalyzed insulin receptor kinase domain fragment KK-1 and inhibits the formation of 3,3 -dityrosine (58).
(2) Regulation of miRNAs. Apigenin regulates miRNAs, which are associated with insulin resistance and glucose homeostasis.
In vitro experiments involving Huh7 cells and in vivo studies investigating miR103 transgenic mice validates apigeninmediated inhibition of the phosphorylation of transactivating response RNA-binding proteins (TRBP). Additionally, miRNAgenerating complexes inhibited, leading to suppression of precursor miRNA103 maturation expressed in liver and fat, resulting in insulin resistance and impaired glucose metabolism and homeostasis (59,60). Thus, apigenin-induced suppression of miRNA103 alleviates glucose intolerance (60).
(3) Upregulation of GLUT4/AMPK signaling. Apigenin extracted from Sophora davidii (Franch.) promotes glucose transporter 4 (GLUT4) expression and activates AMPK phosphorylation in L6 cells and insulin target tissues in KK-Ay mice (61). In insulin target tissues such as liver and fat, the upregulation of GLUT4 and the activation of AMPK facilitates glucose utilization to ameliorate insulin resistance (62,63).
(4) Inhibition of α-amylase. Several studies reported that apigenin decreases the inhibition of α-amylase in Kunming mice, thus reducing the digestion of dietary carbohydrates (64). The digestive enzyme α-amylase hydrolyzes dietary carbohydrates into disaccharides and polysaccharides (65). Inhibition of the digestion of dietary carbohydrates delays glucose absorption and blocks the progression of T2DM. Therefore, the inhibition of α-amylase by apigenin ameliorates T2DM (64). Oxidative stress also triggers β-cell dysfunction, impaired glucose tolerance, and insulin resistance (66). The production of reactive oxygen species (ROS) in oxidative stress exacerbates the progression of T2DM and related complications. Apigenin treatment mitigates oxidative stress and intracellular ROS production via following mechanisms: (1) Decreased ROS production. Apigenin pre-treatment of streptozocin (STZ)-treated RINm5F pancreatic β cells ameliorates STZ-induced intracellular ROS production, as well as DNA damage, lipid peroxidation, and apoptosis. Apigenin pretreatment upregulates the expression of antioxidant enzymes, such as superoxide dismutase (SOD), catalase (CAT), and glutathione peroxidase (GSH-Px) in RINm5F pancreatic β cells and diabetic rats (67). SOD catalyzes the conversion of superoxide radicals (O 2− ) to molecular oxygen (O 2 ) and hydrogen peroxide (H 2 O 2 ), resulting in a protective effect against ROS in cells (68). Catalase is a highly specific enzyme that catalyzes the decomposition of hydrogen peroxide into water and molecular oxygen (69). GSH-Px is a cytosolic enzyme that catalyzes the reduction of hydrogen peroxide and lipid peroxides by glutathione, releasing water, oxygen, and alcohol (70). These three enzymes are indispensable in defending against free radicals (71).
(2) Inhibition of AGE. Apigenin inhibits the formation of advanced glycation end products (AGEs) and thereby alleviates oxidative stress (72). AGE-mediated damage leads to altered protein structure and functions via cross-linking between molecules via the receptor for AGEs (RAGE). AGEs increase ROS formation and damage anti-oxidant systems (73). Apigenin treatment of human blood plasma proteins in vitro reduced the levels of AGEs (72).
(3) Regulation of Keap1-Nrf2 signaling. The anti-oxidant function of apigenin is mediated via the Kelch-like ECHassociated protein 1 (Keap1)-Nrf2 axis targeting liver tissues to alleviate oxidative stress (74). Nrf2 is a primary transcription factor interacting with the anti-oxidant response element (ARE) to regulate antioxidant protein expression. Keap1 is the specific repressor of Nrf2, which acts as an adaptor protein of the Cullin3based ubiquitin E3 ligase complex to facilitate the ubiquitination and subsequent proteolysis of Nrf2 (74), acting as a sensor of oxidative stress (75,76). Apigenin occupies the Nrf2-binding site to prevent the binding between Keap1 and Nrf2 and thereby promotes nuclear translocation of Nrf2, thus facilitating its antioxidant function (74).
Currently, the apigenin-mediated regulation of blood glucose homeostasis can be summarized as follows: regulating the key enzymes and improving oxidative stress as well as inflammation. A detailed summary of the studies discussed above and the proposed mechanisms of apigenin-mediated effects in diabetes are presented in Table 2. Dosages and duration of apigenin treatments in vivo and in vitro are also listed in Table 2.

Protective Role of Apigenin in Hypertension
Hypertension plays a central role in cardiometabolic diseases (79), which is prevalent in almost 80% of patients with metabolic syndrome (80). Recent studies reported that apigenin improves hypertension via attenuation of oxidative stress and recovery of mitochondrial dysfunction.
Apigenin plays a protective role in hypertension by alleviating oxidative stress. Apigenin can significantly restore normal blood pressure and reverse renal damage in cyclosporineinduced hypertensive Sprague-Dawley rats by decreasing lipid hydroperoxides and increasing anti-oxidant levels (81). Apigenin also controls elevated blood pressure in N-nitro-L-arginine methylester-induced hypertensive Sprague-Dawley rats by improving NO bioavailability, attenuating oxidative stress, and reducing vascular damage (82).
Apigenin also regulates pulmonary hypertension (PH). Mitochondrial dysfunction plays a vital role in PH, it may lead to the imbalance of ion homeostasis and downregulation of enzymes in apoptosis (83). Apigenin activates mitochondriadependent apoptosis via hypoxia-inducible factor 1α (HIF-1α)-KV1.5 channel pathway. The inhibition of HIF-1α by apigenin upregulates the expression of KV1.5 channels to restore mitochondrial function, thereby attenuating PH (84).
Apigenin has also been reported to diminish the complications induced by hypertension, such as renal damage and fibrosis due to abnormal collagen accumulation in kidneys (85). Apigenin significantly attenuated hypertension and renal fibrosis in deoxycorticosterone acetate (DOCA)-salt-induced hypertensive rats (86). Apigenin activates transient receptor potential vanilloid 4 (TRPV4), a non-selective cation channel widely expressed in the kidney. Ca 2+ influx is then promoted in vascular endothelium and smooth muscle to induce vasodilation (87) and activation of the AMPK/SIRT1 signaling pathway to inhibit the TGF-β1 and Smad-2/3 signaling pathway (Sma and Mad proteins from Caenorhabditis elegans and Drosophila, respectively). This inhibition stimulates cellular transformation into fibroblasts and increases the synthesis of matrix proteins to induce renal fibrosis (86,88,89). Thus, apigenin alleviates renal fibrosis and structural and functional damage.
Current evidence suggests that apigenin decreases blood pressure mainly via improved NO bioactivity and oxidative stress, regulation of apoptosis-related mitochondrial genes and promotion of vasodilation in vascular endothelium. Experimental studies and mechanisms of action involving apigenin in hypertension are listed in Table 3. Experimental dosages and durations of apigenin treatment in vivo and in vitro are listed in Table 3.

Protective Role of Apigenin in Cardiovascular Diseases
Apigenin prevents cardiovascular diseases via antioxidant and anti-apoptotic mechanisms in vascular endothelial cells and cardiomyocytes.
Vascular endothelial dysfunction is a major mediator in cardiovascular diseases (90). Abnormal glucose metabolism He et al. (84) In vitro experiment HBZY-1 cells, M1CCD cells and oxidative stress in vascular endothelial cells may lead to vascular endothelial dysfunction. Several studies have discussed the ameliorative effect of apigenin on endothelial dysfunction. First, apigenin increases apelin expression to rescue endothelial dysfunction. Apelin is an endogenous ligand for the G-proteincoupled APJ receptor expressed in the cardiovascular system. It increases glucose uptake and SOD activity, reversing the impaired glucose metabolism and homeostasis and the severe oxidative stress in human endothelial cells (91)(92)(93)(94). Oxidative stress in endothelial cells leads to endothelial dysfunction and angiogenesis (92). The suppression of apelin in human endothelial cells can be reversed by apigenin treatment. Second, apigenin inhibits NF-κB-associated signaling pathways and suppresses intercellular adhesion molecule-1 (ICAM-1) expression in vascular endothelial dysfunction. ICAM-1 is a cell surface receptor that binds lymphocyte functionassociated antigen 1 (LFA-1), mediating the interaction between keratinocytes and leukocytes (95). ICAM-1 plays an essential role in controlling abnormal inflammatory infiltration, adhesion, and migration (96). Apigenin inhibits NF-κB activation to improve NO production and SOD activity in endothelial cells and suppress ICAM-1 expression in human vascular endothelial cells (HUVECs) (97). Third, apigenin inactivates the PI3K (phosphoinositide-3-kinase) Akt (protein kinase B) axis in HUVECs during vascular endothelial dysfunction. The PI3K/Akt axis is an essential pathway in the pathogenesis of cardiovascular complications in T2DM (98). Apigenin treatment inhibited the phosphorylation of Akt-residues Ser473 and Thr308 to prevent vascular endothelial dysfunction (99). Finally, apigenin decreased ROS and improved NO levels to alleviate vascular endothelial dysfunction induced by mitochondriadependent apoptosis via inhibition of protein kinase C βII (PKCβII) phosphorylation. PKCβII promotes oxidative stress, ROS production and mitochondria-dependent apoptosis in vascular endothelial dysfunction (100,101). Apigenin treatment upregulated the expression of the anti-apoptotic gene, B-cell lymphoma-2 (Bcl-2), while the pro-apoptotic gene, Bcl-2 associated X (Bax), was downregulated, resulting in attenuation of mitochondria-dependent apoptosis in endothelial cells (102). Further, cardiac hypertrophy is another manifestation of cardiovascular diseases (103). Current evidence suggests that abnormal glycolipid metabolism and overexpression of HIF-1α in cardiac cells causes cardiac hypertrophy (104). In vivo and in vitro experimental data suggest that apigenin alleviates cardiac hypertrophy via suppression of HIF-1α, thereby reversing the expression of PPARα/γ and target genes including glycerol-3-phosphate acyltransferase (GPAT), glucose transporter 4 (GLUT-4), carnitine palmitoyltransferase 1 (CPT-1) and pyruvate dehydrogenase kinase 4 (PDK-4). Downregulation of GLUT4 and upregulation of PDK-4 can inhibit excessive glucose intake and oxidation, preventing abnormal glucose metabolism. Downregulation of GPAT and upregulation of CPT-1 decreases the rate of triglyceride synthesis and augments fatty acid oxidation, thereby improving lipid metabolism (105,106). Thus, the hypoxic myocardial energy utilization (107)(108)(109)(110) can be reversed. Ren et al. (97) In summary, apigenin can ameliorate cardiovascular diseases via reduction of oxidative stress and mitochondria-dependent apoptosis in vascular endothelial cells as well as regulation of glucose and lipid metabolism in cardiomyocytes. The experimental studies and mechanisms of action involving apigenin in hypertension are presented in Table 4. Experimental dosages and duration of apigenin treatment in vivo and in vitro are also listed in Table 4.

Apigenin Analogs and Their Effects on Alleviating Cardiometabolic Diseases
Apigenin analogs are derived from the basic flavonoid skeleton via hydroxyl group substitution, glycosylation, hydroxylation, and methylation (111,112). In plants, apigenin is stored as glycosides such as apigenin 7-O-apioglucoside in celery and parsley (113,114) and apigenin 8-C-glucoside isolated from bamboo leaves (115). Several apigenin analogs carry the basic flavonoid skeleton similar to apigenin and exhibit biological activity in cardiometabolic diseases.

POTENTIAL SIGNALING PATHWAYS MEDIATED BY APIGENIN FOR AMELIORATION OF CARDIOMETABOLIC DISEASE IN DIFFERENT CELL TYPES
In summary, the potential signaling pathways mediated by apigenin resulting in alleviation of cardiometabolic diseases in different cell types are illustrated in Figures 2A-E. Apigenin alleviates cardiometabolic diseases mainly by regulating glycolipid metabolism, oxidative stress, and oxidative stressinduced inflammation and apoptosis. Notably, apigenin plays contrasting roles in different types of cells. Apigenin acts as an agonist of PPARγ in adipose tissue macrophages. PPARγ binds to p65 to inhibit nuclear translocation to block NF-κB signaling pathway resulting in attenuation of inflammation (Figure 2A) (33). However, apigenin inhibits PPARγ expression in adipocytes, hepatocytes, and cardiomyocytes by acting as an antagonist (Figures 2B-D).

Apigenin in Adipocytes
In adipocytes, apigenin acts as a functional regulator of lipid metabolism to reduce fat accumulation. Apigenin downregulates PPARγ expression by inhibiting the STAT3/CD36 axis (16) and the activation of AMPK (17). Apigenin directly induces p65 phosphorylation to prevent its nuclear translocation to ensure continued inhibition of NF-κB signaling in the absence of PPARγ as a mediator (25,35).

Apigenin in Hepatocytes
Since the liver is an important site of energy metabolism, apigenin is a potential mediator of glycolipid metabolism in hepatocytes. Apigenin acts as a PPARγ antagonist via direct activation of Nrf2 and indirect activation of Nrf2 via the Keap1-Nrf2 pathway (126). Additionally, apigenin activates AMPK to inhibit SREBP-1 and SPEBP-2 to regulate hepatic fatty acid oxidation and cholesterol synthesis (28).

Apigenin in Cardiomyocytes
Apigenin treatment of cardiomyocytes regulates glucose and lipid metabolism to maintain normal cellular function. HIF-1α activation via apigenin regulates the PPAR family, leading to the appropriate regulation of downstream target genes related to glycolipid metabolism. Apigenin suppresses PPARγ expression via the activation of HIF-1α as an antagonist (105,106). Meanwhile, the upregulation of HIF-1α following apigenin treatment increases PPARα expression (105,106).

Apigenin in Endothelial Cells
Apigenin plays a protective role in vascular endothelial dysfunction by regulating several signaling pathways in endothelial cells to alleviate oxidative stress, inflammation, and mitochondria-dependent apoptosis. The inhibition of the NF-κB signaling pathway with apigenin treatment also ameliorates inflammatory response in endothelial cells and increases NO production (97). NF-κB inhibition also suppresses the expression of ICAM-1 to improve abnormal inflammatory adhesion, migration, and infiltration, resulting in the alleviation of vascular endothelial dysfunction (97). In addition to NF-κB pathways, apigenin also activates the PI3K-Akt pathway and inhibits PKCβII activation by reducing oxidative stress and oxidative stress-related apoptosis in mitochondria (102). The expression of anti-apoptotic gene, Bcl-2, and the pro-apoptotic gene, Bax, in these two pathways reduces abnormal apoptosis. The activation of the PI3K-Akt pathway also promotes eNOS activity to restore NO levels and thereby attenuates oxidative stress (102). Apigenin treatment also mitigates angiogenesis induced by inflammation. HIF-1α inhibition by apigenin directly reduces the expression of vascular endothelial growth factor (VEGF) in angiogenesis, thus alleviating the angiogenesis induced by vascular dysfunction (127).

CONCLUSION AND PERSPECTIVES
A review of studies investigating apigenin suggests critical biological mechanisms, including reducing oxidative stress and oxidative stress-induced inflammation and apoptosis, and improving glycolipid metabolism. Figure 3 summarizes the potential signaling pathways of apigenin underlying the protection against cardiometabolic diseases.
The molecular structure of apigenin suggests poor water solubility, chemical instability and moderate permeability, which prevent maximum bioavailability. Therefore, new delivery and design strategies have been formulated including the development of apigenin glycosides and acylated derivatives to enhance water solubility (128,129). Apigenin-AuNP complex can be developed at room temperature at pH 10 to enhance the stability of apigenin in the body (130). Nano-apigenin using poly (lactic-co-glycolide) (PLGA) can also improve the bioactivity of apigenin (131). Pharmacokinetic and pharmacodynamic profiles of apigenin in rats and mice have been studied. The peak plasma concentration C max and the time to reach the peak plasma concentration T max were 1.07 ng/mL and 1 h, respectively, and the area under the concentration-time curve (AUC 0−24 ) was 3.9 ng h/mL in mice (132). However, the bioavailability of apigenin in humans is still unknown. Further studies are needed to confirm the bioavailability and safety profile in humans.
In summary, the extensive review and validation of in vitro and in vivo evidence suggests that apigenin is a natural compound that can be used to protect against cardiometabolic diseases. Environment-wide association studies (EWAS) also indicate that apigenin is one of the protective factors in cardiovascular diseases at the population level (133). Further studies are required to establish the optimum dose of apigenin in alleviating cardiometabolic diseases in humans, developing a novel approach for clinical management of the disease.

AUTHOR CONTRIBUTIONS
XL designed this review, helped with writing and revising of the manuscript, and provided critical feedback. YX contributed to collecting and screening the literature, as well as summarizing the data, and then composed and revised the manuscript. HW reviewed the manuscript. All authors were involved in final approval of the submitted version.