Serial Lipocalin 2 and Oncostatin M levels reflect inflammation status and treatment response in axial spondyloarthritis

Background Informative serum biomarkers for monitoring inflammatory activity and treatment responses in axial spondyloarthritis (axSpA) are lacking. We assessed whether Lipocalin 2 (LCN2) and Oncostatin M (OSM), both having roles in inflammation and bone remodeling, may accurately reflect chronic joint inflammation and treatment response in axSpA. Previous reports in animal models showed involvement of LCN2 and OSM in joint/gut inflammation. We asked whether they also play a role in human axSpA. Methods We analyzed a longitudinal observational axSpA cohort (286 patients) with yearly clinical assessments and concurrent measurements of serum LCN2 and OSM (1204 serum samples) for a mean of 4 years. Biomarker levels were correlated with MRI scoring and treatment response. Results Persistent and transient elevation of LCN2 and OSM were observed in axSpA patients. Persistent elevation of LCN2 or OSM, but not CRP, correlated with sacroiliac joint (SIJ) MRI SPARCC scores (Pearson's correlation p = 0.0005 and 0.005 for LCN2 and OSM respectively), suggesting that LCN2/OSM outperforms CRP as reflective of SIJ inflammation. We observed both concordant and discordant patterns of LCN2 and OSM in relationship to back pain, the cardinal clinical symptom in axSpA. Twenty-six percent (73/286) of the patients remained both clinically and serologically active (CASA). Sixty percent (173/286) of the patients became clinically quiescent, with back pain resolved, but 53% (92/173) of them were serologically active (CQSA), indicating that pain control may not indicate control of joint inflammation, as reflected by positive MRI imaging of SIJ. With respect to treatment responses, transient elevation of LCN2 or OSM over time was predictive of better response to all treatments. Conclusion In axSpA, persistent LCN2 and/or OSM elevation reflects chronic SIJ inflammation and suboptimal treatment response. In our cohort, half of the currently deemed clinically quiescent patients with back pain resolved continued to demonstrate chronic joint inflammation. LCN2 and OSM profiling outperforms CRP as a predictive measure and provides an objective assessment of chronic local inflammation in axSpA patients

Lipocalin 2 links inflammation and ankylosis in the clinical overlap of inflammatory bowel disease (IBD) and ankylosing spondylitis (AS)

Abstract Background Little is known about the mechanisms underlying the clinical overlap between gut inflammation and joint ankylosis, as exemplified by the concurrence of inflammatory bowel diseases (IBD) and ankylosing spondylitis (AS). As dysbiosis may serve as a common contributor, the anti-microbial pleiotropic factor lipocalin 2 could be a potential mediator due to its roles in inflammation and bone homeostasis. Methods Baseline colonic pathology was conducted in the ank/ank mouse model. Serum lipocalin 2 was analyzed by ELISA, in ank/ank mutants versus C3FeB6-A/Aw-jwt/wt, in patients with concurrent AS-IBD, AS alone, IBD alone, or mechanical back pain, and in healthy controls. In the ank/ank mouse model, the expression of nuclear receptor peroxisome proliferator-activated receptor gamma (PPARγ) was examined by real-time PCR. Intraperitoneal injection was done with the PPARγ agonist rosiglitazone or antagonist bisphenol A diglycidyl ether for four consecutive days. Serum levels of lipocalin 2 were examined on the sixth day. Results This study showed that the ank/ank mice with fully fused spines had concurrent colonic inflammation. By first using the ank/ank mouse model with progressive ankylosis and subclinical colonic inflammation, confirmed in patients with concurrent AS and IBD, elevated circulating lipocalin 2 levels were associated with the coexisting ankylosis and gut inflammation. The intracellular pathway of lipocalin 2 was further investigated with the ank/ank mouse model involving PPARγ. Colonic expression of PPARγ was negatively associated with the degree of gut inflammation. The PPARγ agonist rosiglitazone treatment significantly upregulated the serum levels of lipocalin 2, suggesting a potential regulatory role of PPARγ in the aberrant expression of lipocalin 2. Conclusions In summary, lipocalin 2 modulated by PPARγ could be a potential pathway involved in concurrent inflammation and ankylosis in AS and IBD

The role of LCN2 and LCN2-MMP9 in spondylitis radiographic development: gender and HLA-B27 status differences

Background Male HLA-B27-positive radiographic-axial spondyloarthritis (r-axSpA) patients are prone to have severe spinal radiographic progression, but the underlying mechanisms are unclear. We recently showed that persistently elevated Lipocalin 2 (LCN2; L) reflects sacroiliac joint (SIJ) inflammation. LCN2 binds to MMP9. Concomitant elevation of L and LCN2-MMP9 (LM) was detected in many inflammatory diseases. We asked whether L and LM play similar roles in r-axSpA pathogenesis. Methods We analyzed 190 axSpA patients (123 radiographic and 67 non-radiographic axSpA) who had no detectable circulating Oncostatin M, to avoid complications due to cross-talk between pathways. L and LM levels from a single blood sample of each patient were measured and were correlated with MRI and modified stoke AS (mSASS) scoring. Association of elevated L (L+) or concurrent L+ and elevated LM (LM+) patterns with B27 status and gender were assessed. Results In L+LM+ axSpA patients, both L and LM levels correlated with MRI SPARCC SIJ scores, but only LM levels correlated with MRI Berlin Spine Scores, suggesting LM is a biomarker for both SIJ and spinal inflammation. Among patients with minimal spinal ankylosis (mSASSS < 10), 65% of male r-axSpA patients are L+LM+, while 30% and 64% of female patients are L+LM+ and L+, respectively, supporting the role of LM with disease progression. In B27+ L+LM+ male patients, both L and LM (but not CRP) levels correlate with mSASSS. B27 positivity and maleness have additive effects on spondylitis progression, suggesting concurrent high L and LM elevations are associated with B27+ male patients having more significant radiographic damage. L+ B27-negative male patients or L+ female patients are more likely to have milder disease. Conclusion L and LM are informative biomarkers for SIJ and spinal inflammation, as well as for ankylosing development in r-axSpA patients. Distinctive L+LM+ or L+ patterns not only could distinguish clinically aggressive vs milder course of disease, respectively, but also provide an explanation for B27-positive male patients being the most susceptible to severe ankylosis

Regulation of hERG and hEAG channels by Src and by SHP-1 tyrosine phosphatase via an ITIM region in the cyclic nucleotide binding domain.

Members of the EAG K(+) channel superfamily (EAG/Kv10.x, ERG/Kv11.x, ELK/Kv12.x subfamilies) are expressed in many cells and tissues. In particular, two prototypes, EAG1/Kv10.1/KCNH1 and ERG1/Kv11.1/KCNH2 contribute to both normal and pathological functions. Proliferation of numerous cancer cells depends on hEAG1, and in some cases, hERG. hERG is best known for contributing to the cardiac action potential, and for numerous channel mutations that underlie 'long-QT syndrome'. Many cells, particularly cancer cells, express Src-family tyrosine kinases and SHP tyrosine phosphatases; and an imbalance in tyrosine phosphorylation can lead to malignancies, autoimmune diseases, and inflammatory disorders. Ion channel contributions to cell functions are governed, to a large degree, by post-translational modulation, especially phosphorylation. However, almost nothing is known about roles of specific tyrosine kinases and phosphatases in regulating K(+) channels in the EAG superfamily. First, we show that tyrosine kinase inhibitor, PP1, and the selective Src inhibitory peptide, Src40-58, reduce the hERG current amplitude, without altering its voltage dependence or kinetics. PP1 similarly reduces the hEAG1 current. Surprisingly, an 'immuno-receptor tyrosine inhibitory motif' (ITIM) is present within the cyclic nucleotide binding domain of all EAG-superfamily members, and is conserved in the human, rat and mouse sequences. When tyrosine phosphorylated, this ITIM directly bound to and activated SHP-1 tyrosine phosphatase (PTP-1C/PTPN6/HCP); the first report that a portion of an ion channel is a binding site and activator of a tyrosine phosphatase. Both hERG and hEAG1 currents were decreased by applying active recombinant SHP-1, and increased by the inhibitory substrate-trapping SHP-1 mutant. Thus, hERG and hEAG1 currents are regulated by activated SHP-1, in a manner opposite to their regulation by Src. Given the widespread distribution of these channels, Src and SHP-1, this work has broad implications in cell signaling that controls survival, proliferation, differentiation, and other ERG1 and EAG1 functions in many cell types

EAG-family members contain a conserved ITIM motif.

<p>Sequence alignments of C-termini for the EAG family of K⁺ channels show each immuno-receptor tyrosine inhibitory motif (ITIM) in bold, with its potentially phosphorylated tyrosine residue in parentheses beside the channel name. Also shown for hERG/Kv11.1/KCHN2 is the sequence of the 15 amino acid peptide made for use in pull-down assays or deleted in the ΔITIM mutant channel (line above the sequence).</p

The hERG current in stably transfected HEK293 cells is reduced by the Src-family tyrosine kinase inhibitor, PP1.

<p><b>A. i.</b> Representative current traces in response to the voltage-clamp protocol shown in the inset. From the holding potential of −80 mV, 3-s long depolarizing steps were applied between −70 and +60 mV (10-mV increments). This opens the channels, but at positive potentials, some inactivation is seen (c→o→i). The voltage was then stepped to −40 mV to remove inactivation (i→o) and monitor outward tail currents. <b>ii.</b> The current amplitude at the end of each 3-sec long depolarizing step (open arrow in Ai) was used to construct a current-versus-voltage (I–V curve) before and after perfusing the hERG blocker, 1 µM E-4031, into the bath. <b>iii.</b> Summary of instantaneous I–V relations from tail currents (closed arrow in Ai) before and after adding E-4031. In all graphs, the values are mean±SEM for the number of cells in parentheses. <b>B.</b> Representative whole-cell currents are shown before and 25 min after adding 10 µM PP1 to the bath. From a holding potential of −80 mV, a 1 s long pulse to +60 mV was used to activate and then inactivate the channels (c→o→i). Then, inactivation was rapidly removed by a 16 ms long pulse to −120 mV (i→o). Instantaneous tail currents (arrow) during test pulses between −110 and +20 mV (10-mV increments) were used to measure the open-channel current-<i>vs</i>-voltage (I–V) relationship (summarized in D) and inactivation kinetics (summarized in E). <b>C.</b> PP1 reduces the hERG current. To monitor time-dependent changes, the instantaneous tail current at +20 mV was repeatedly measured in control cells and in separate cells exposed to 10 µM PP1. Each current was normalized to its initial value (measured at 3–4 min after starting the recording). Significant differences between control and PP1-treated cells are indicated as *<i>p</i><0.05; **<i>p</i><0.01. <b>D.</b> Open-channel I–V relations from instantaneous tail currents (as in panel B) were recorded at 25 min in control cells and in separate cells exposed to 10 µM PP1. For each cell, the current was normalized to cell capacitance to yield current density (pA/pF). <b>E.</b> The time constant of inactivation (τ) was measured after 25 min in control cells or with 10 µM PP1 in the bath. That is, for test pulses between +20 and −20 mV, the decay of the tail current (o→i; as in panel B) was fitted to a mono-exponential function: <i>I_t</i> = <i>AMP</i>*exp(−<i>t/τ</i>), where <i>I_t</i> is the outward current at time <i>t</i> and <i>AMP</i> is the initial current amplitude. Each fit was begun at the time indicated by the arrow in panel B.</p

Src tyrosine kinase regulates the hERG current.

<p>Currents were compared between cells recorded with pipette solutions containing saline (to control for spontaneous changes), 100 µg/mL of a Src-inhibitory peptide (Src40-58), or 100 µg/mL of a scrambled version of the same peptide (Src40-58s). For each cell, the voltage protocol was repeated at 0, 10, 20 and 30 min of recording; and the representative current traces (upper panels) are from 2 control cells and 2 treated cells, all at 30 min. <b>A.</b> The voltage protocol was similar to <a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0090024#pone-0090024-g001" target="_blank">Fig. 1B</a>; i.e., from a holding potential of −80 mV, a 1 s long pulse to +60 mV was applied to activate the channels. Then inactivation was removed by a 16 ms long pulse to −120 mV, and the maximal outward tail current was measured during a test pulse to +20 mV. The lower panel summarizes the time course, measured from the maximal tail current, normalized to its initial value at 3–4 min after establishing a recording. <b>B.</b> Following a 6 s long pre-pulse to +20 mV to activate the channels, the maximal inward tail current was measured during the test pulse to −120 mV (similar protocol to <a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0090024#pone-0090024-g002" target="_blank">Fig. 2A</a>). The summarized time course in the lower panel was constructed as in panel A. Significant differences for Src40-58 are shown compared with controls (*<i>p</i><0.05) or the scrambled peptide (^#<i>p</i><0.05).</p

The hEAG1/KCNH1 current is regulated by SHP-1 tyrosine phosphatase.

<p>The voltage protocols and normalization procedures were the same as in <a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0090024#pone-0090024-g004" target="_blank">Fig. 4</a> (only traces at +60 mV are shown). <b>A.</b> Time-dependent effects of adding 100 µg/mL recombinant wild-type SHP-1 protein to the pipette solution. Inset: representative currents from two cells, recorded 30 min after establishing the whole-cell configuration. For each cell, the peak current at +60 mV was repeatedly measured and normalized to its initial value (3–4 min after establishing a recording) for the number of cells indicated (*<i>p</i><0.05). <b>B.</b> Comparison of cells recorded with control pipette solution with those containing a 3∶1 mixture of the inactive substrate-trapping SHP-1 mutant (SHP-1 C453S; 150 µg/mL) and active wild-type SHP-1 (50 µg/mL). Inset: representative currents recorded 30 min after establishing the whole-cell configuration. The normalized peak currents at +60 mV are shown; *<i>p</i><0.05.</p

PP1 reduces the hERG window current without apparently affecting the voltage dependence or kinetics.

<p>Whole-cell hERG currents in stably transfected HEK293 cells, with representative current traces in response to the voltage-clamp protocols shown as insets. For all panels, control recordings at 25 min are compared with 25 min after adding 10 µM PP1 to the bath. <b>A.</b> The voltage dependence of activation was monitored by varying the 6 s long pre-pulse between −100 and +40 mV (20 mV increments) (c→o→i). Then, the maximal inward current was measured during a test pulse to −120 mV, which rapidly relieved inactivation (i→o) and revealed transient hERG currents before the channels slowly closed (o→c). The peak amplitudes were used to construct activation-<i>vs</i>-voltage curves (summarized in panel E). <b>B.</b> A single protocol was used to monitor the voltage dependence of inactivation, and the time courses of recovery from inactivation and channel closing. From a holding potential of −80 mV, a 1 s-long pre-pulse to +60 mV activated and then inactivated the channels (c→o→i). Then, test pulses were applied between −120 and +20 mV (10 mV increments). The rapid increase in inward currents reflects recovery from inactivation (i→o; summarized in C), and the subsequent slow decline reflects closing (o→c; summarized in D). The maximal current at each test voltage was used to construct inactivation-<i>vs</i>-voltage curves (summarized in panel E). <b>C, D.</b> Time constants (τ) of recovery from inactivation (C) and closing (D) are summarized. For test potentials between −120 and −90 mV, the currents (as in panel B) were fitted to a dual exponential (rising and falling) function: <i>I_t</i> = <i>I_o</i>+(<i>AMP_r</i>*exp(−<i>t/τ_r</i>)+<i>AMP_d</i>*exp(−<i>t/τ_d</i>)). The values are presented as mean±SEM for the number of cells in parentheses. <b>E.</b> Activation-<i>vs</i>-voltage curves were obtained by fitting the peak currents (examples in panel A) to a Boltzmann equation: <i>G/G_max</i> = 1/(1+exp[(<i>V_m</i>−<i>V_1/2</i>)/<i>k</i>]), where <i>V_m</i> is membrane potential, <i>V_1/2</i> is the potential at which <i>G/G_max</i> = 0.5, and <i>k</i> is the slope factor. The degree of inactivation as a function of voltage (data as in panel B) was calculated from: <i>I</i>/(<i>G</i>_slope(<i>V_m</i>−<i>E</i>_K)), where I is the peak current at each voltage, <i>V_m</i> is the test potential, <i>E</i>_K is the K⁺ Nernst potential, and <i>G</i>_slope is the maximal slope conductance, calculated from a linear fit to the I–V relation between −120 and −90 mV (not shown). <b>F.</b> The window current was calculated by multiplying the results of each fitted equation (panel E) by the maximal whole-cell conductance and dividing by the driving force (V_m−E_K).</p