Letter | Published:

Adult hippocampal neurogenesis is abundant in neurologically healthy subjects and drops sharply in patients with Alzheimer’s disease

Nature Medicinevolume 25pages554560 (2019) | Download Citation

Abstract

The hippocampus is one of the most affected areas in Alzheimer’s disease (AD)1. Moreover, this structure hosts one of the most unique phenomena of the adult mammalian brain, namely, the addition of new neurons throughout life2. This process, called adult hippocampal neurogenesis (AHN), confers an unparalleled degree of plasticity to the entire hippocampal circuitry3,4. Nonetheless, direct evidence of AHN in humans has remained elusive. Thus, determining whether new neurons are continuously incorporated into the human dentate gyrus (DG) during physiological and pathological aging is a crucial question with outstanding therapeutic potential. By combining human brain samples obtained under tightly controlled conditions and state-of-the-art tissue processing methods, we identified thousands of immature neurons in the DG of neurologically healthy human subjects up to the ninth decade of life. These neurons exhibited variable degrees of maturation along differentiation stages of AHN. In sharp contrast, the number and maturation of these neurons progressively declined as AD advanced. These results demonstrate the persistence of AHN during both physiological and pathological aging in humans and provide evidence for impaired neurogenesis as a potentially relevant mechanism underlying memory deficits in AD that might be amenable to novel therapeutic strategies.

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Data availability

The data that support the findings of this study are available from the corresponding author upon reasonable request. All requests for raw and analyzed data will be promptly reviewed by the Center for Networked Biomedical Research on Neurodegenerative Diseases (CIBERNED) to determine whether the request is subject to any intellectual property or confidentiality obligations. Any materials that can be shared will be released via a material transfer agreement.

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Acknowledgements

The authors would like to thank the patients and their families for generously donating brain samples. Moreover, they would like to thank I. Rodal for help with human sample extraction and processing, E. García, R. Cuadros and the confocal microscopy facility of the CBMSO for technical assistance, and P. Moreno for help with illustration design. A number of human samples were generously provided by the Biobanco del Hospital Universitario Reina Sofia (Córdoba, Spain). The authors are grateful to R. Sánchez for providing some of these samples. They would also like to thank J. Gleeson (University of California, San Diego) for providing an anti-DCX antibody. This study was supported by the following: the Spanish Ministry of Economy and Competitiveness (SAF-2017-82185-R and RYC-2015-171899, M.L.-M.; SAF-2014-53040-P, J.Á.); the Alzheimer’s Association (2015-NIRG-340709 and AARG-17-528125, M.L.-M.); the Association for Frontotemporal Degeneration (2016 Basic Science Pilot Grant Award, M.L.-M.); the Comunidad de Madrid (PEJD-2017-PRE/BMD-3439, M.L.-M.); and the Center for Networked Biomedical Research on Neurodegenerative Diseases (CIBERNED, Spain, J.Á.). Institutional grants from the Fundación Ramón Areces and the Banco de Santander to CBMSO are also acknowledged. The salary of E.P.M.-J. was supported by a Comunidad de Madrid researcher contract (PEJD-2017-PRE/BMD-3439). The salary of J.T.-R. was supported by a Universidad Autónoma de Madrid doctoral fellowship (FPI-UAM 2017 program).

Author information

Author notes

  1. These authors contributed equally: E. P. Moreno-Jiménez, M. Flor-García, J. Terreros-Roncal.

Affiliations

  1. Department of Molecular Neuropathology, Centro de Biología Molecular ‘Severo Ochoa’, CBMSO, CSIC-UAM, Madrid, Spain

    • Elena P. Moreno-Jiménez
    • , Miguel Flor-García
    • , Julia Terreros-Roncal
    • , Noemí Pallas-Bazarra
    • , Jesús Ávila
    •  & María Llorens-Martín
  2. Department of Molecular Biology, Faculty of Sciences, Universidad Autónoma de Madrid, Madrid, Spain

    • Elena P. Moreno-Jiménez
    • , Miguel Flor-García
    • , Julia Terreros-Roncal
    •  & María Llorens-Martín
  3. Center for Networked Biomedical Research on Neurodegenerative Diseases (CIBERNED), Madrid, Spain

    • Elena P. Moreno-Jiménez
    • , Miguel Flor-García
    • , Julia Terreros-Roncal
    • , Noemí Pallas-Bazarra
    • , Jesús Ávila
    •  & María Llorens-Martín
  4. Neuropathology Department, CIEN Foundation, Madrid, Spain

    • Alberto Rábano
  5. Universidad Europea de Madrid, Faculty of Biomedical and Health Sciences, Madrid, Spain

    • Fabio Cafini

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Contributions

E.P.M.-J., M.F.-G., J.T.-R., and M.L.-M. designed and conceived the experiments. A.R. provided materials and performed autopsies. M.L.-M. sectioned human samples. E.P.M.-J., M.F.-G., J.T.-R., A.R., F.C., N.P.-B., and M.L.-M. performed the experiments and acquired confocal images. E.P.M.-J., F.C., and M.L.-M. performed cell counts and analyzed the data. M.L.-M. wrote the manuscript. J.Á. and M.L.-M. obtained funding. All authors critically discussed the data and revised the final version of the manuscript.

Competing interests

The authors declare no competing interests.

Corresponding author

Correspondence to María Llorens-Martín.

Extended data

  1. Extended Data Fig. 1 Control subjects included in this work.

    a, Epidemiological data for the 13 neurologically healthy control subjects included in this study. Age, sex, PMD, cause of death, type of fixative, and fixation conditions are indicated. A tissue block from each individual was fixed for 24 h in 4% PFA at 4 °C. Moreover, additional blocks of tissue from control 2 and control 4 were fixed in formalin for 6 months. Cases indicated with an exclamation mark were included in control fixation experiments, and additional blocks of tissue were therefore fixed for 1, 2, 6, 12, and 48 h in 4% PFA at 4 °C. b, Correlation between the age of subjects and the density of DCX+ cells. There is a negative correlation between these parameters (two-sided Pearson’s correlation, r = −0.5842, P = 0.036). c, Correlation between PMD and the density of DCX+ cells (two-sided Pearson’s correlation, r = 0.1027, P = 0.7508). d, Density of DCX+ cells in male and female subjects (Mann–Whitney test, U2,13 = 17.00, P = 0.9399). The graph represents mean values ± s.e.m. e, Correlation between age of the subject and the density of NeuN+ neurons in the GCL (two-sided Pearson’s correlation, r = 0.07694, P = 0.8122). f, Correlation between PMD and the density of NeuN+ neurons in the GCL (two-sided Pearson’s correlation, r = 0.4221, P = 0.1717). g, Influence of sex on the density of NeuN+ neurons in the GCL (Mann–Whitney test, U2,13 = 14.00, P = 0.8081). The graph represents mean values ± s.e.m. h, Correlation between age of the subject and the percentage of DCX+ cells that are CR+ (two-sided Pearson’s correlation, r = −0.3082, P = 0.3565). i, Correlation between PMD and the percentage of DCX+ cells that are CR+ (two-sided Pearson’s correlation, r = 0.03674, P = 0.9146). j, Influence of sex on the percentage of DCX+ cells that are CR+ (Mann–Whitney test, U2,13 = 14.00, P = 0.8081). The graph represents mean values ± s.e.m. k, Correlation between age of the subject and the percentage of DCX+ cells that are CB+ (two-sided Pearson’s correlation, r = −0.2768, P = 0.4710). l, Correlation between PMD and the percentage of DCX+ cells that are CB+ (two-sided Pearson’s correlation, r = −0.1907, P = 0.6230). m, Influence of sex on the percentage of DCX+ cells that are CB+ (Mann–Whitney test, U2,13 = 5.00, P = 0.1714). The graph represents mean values ± s.e.m. n, Correlation between age of the subject and the percentage of DCX+ cells that are PSA-NCAM+ (two-sided Pearson’s correlation, r = −0.1481, P = 0.6291). o, Correlation between PMD and the percentage of DCX+ cells that are PSA-NCAM+ (two-sided Pearson’s correlation, r = −0.3241, P = 0.2526). p, Influence of sex on the percentage of DCX+ cells that are PSA-NCAM+ (Mann–Whitney test, U2,13 = 9.00, P = 0.2828). The graph represents mean values ± s.e.m. q, Correlation between age of the subject and the percentage of DCX+ cells that are PH3+ (two-sided Pearson’s correlation, r = 0.4012, P = 0.1742). r, Correlation between PMD and the percentage of DCX+ cells that are PH3+ (two-sided Pearson’s correlation, r = 0.00116, P = 0.9970). s, Influence of sex on the percentage of DCX+ cells that are PH3+ (Mann–Whitney test, U2,13 = 15.50, P = 0.7573). The graph represents mean values ± s.e.m. In bs, n = 13 control subjects. Twenty stacks of images were obtained for each subject. In colocalization analyses, at least 100 cells per subject were analyzed for each cell marker. *0.05 > P ≥ 0.01 in two-sided Pearson’s correlation.

  2. Extended Data Fig. 2 Influence of tissue fixation on the visualization of human adult hippocampal neurogenesis.

    ak, Effects of fixation time. a, Schematic diagram of the experimental design. Briefly, a hippocampal fragment from four control subjects (61–87 years of age; Extended Data Fig. 1) was dissected and cut into 0.5-cm-thick blocks along the rostro–caudal axis of this structure. Each block was fixed for a different period of time in 4% PFA at 4 °C. bf, Influence of fixation time on the visualization of DCX staining in the human DG. b, Number of DCX+ cells detected after different fixation times (one-way ANOVA, F8,32 = 23.90, P < 0.0001). The graph represents mean values ± s.e.m. cf, Representative images of DCX+ cells, showing the differences in signal quality and background intensity caused by different fixation times. Weak tissue preservation, high-intensity background, and low intensity and specificity of the signal were observed after 1 h of fixation in 4% PFA. In contrast, 2–12 h of fixation in 4% PFA rendered a good signal/background ratio, although tissue robustness was minimal. In contrast, 24 and 48 h of fixation almost abolished DCX detection and substantially increased nonspecific background. Thus, given the marked decay in DCX signal after 12 h of fixation, samples fixed for 24 and 48 h were subjected to aldehyde elimination with 0.5% NaBH4 and to a HC-AR protocol, which revealed the presence of thousands of DCX+ cells in the adult human DG. Moreover, NaBH4 and HC-AR allowed unambiguous identification of neuronal characteristics in these cells. gk, Influence of fixation time on the visualization of PSA-NCAM staining in the human DG. g, Number of PSA-NCAM+ cells detected (one-way ANOVA, F8,32 = 16.54, P < 0.0001). The graph represents mean values ± s.e.m. hk, Representative images of PSA-NCAM staining in each fixation condition tested. Fixation times longer than 12 h impeded PSA-NCAM detection, and incubation with NaBH4 + HC-AR was necessary to allow stereological estimation of the density of PSA-NCAM+ cells. These histological pretreatments allowed identification of thousands of PSA-NCAM+ cells in the DG. In ak, n = 4 control subjects. 10–20 measurements were performed for each subject. lv, Effects of formalin fixation on DCX and PSA-NCAM detection in the DG. l, Experimental design. Briefly, the hippocampi of two control subjects (control 2 and control 4; Extended Data Fig. 1) were dissected, and two 0.5-cm-thick blocks per subject were obtained. One of these blocks was fixed in 4% PFA for 24 h, whereas the other block was stored in 3.7% formalin for 6 months. mp, Representative images of DCX staining in the different experimental conditions. q, Number of DCX+ cells detected in samples fixed either in 4% PFA for 24 h or in 3.7% formalin for 6 months, with or without NaBH4 + HC-AR pretreatment. The graph represents mean values. NaBH4 + HC-AR steps increase the number of DCX+ cells detected in samples fixed with both PFA and formalin. However, neither the intense background nor the marked decay in DCX signal caused by formalin fixation was counteracted by NaBH4 + HC-AR. ru, Representative images of PSA-NCAM staining in the different experimental conditions. v, Number of PSA-NCAM+ cells detected in samples fixed either in 4% PFA for 24 h or in 3.7% formalin for 6 months, with or without NaBH4 + HC-AR. The graph represents mean values. NaBH4 + HC-AR increased the number of PSA-NCAM+ cells detected in samples fixed with PFA. However, formalin fixation abolished the PSA-NCAM signal, and this effect was not prevented by NaBH4 + HC-AR. In lv, n = 2 control subjects. 10–20 measurements were performed for each subject. Yellow scale bars, 50 μm. Green triangles indicate DCX+ cells; red triangles indicate PSA-NCAM+ cells. In b and g, asterisks indicate significant differences with respect to the samples fixed for 1 h in Tukey post hoc comparisons. *0.05 > P ≥ 0.01; **0.01 > P ≥ 0.001; ***P < 0.001.

  3. Extended Data Fig. 3 Identification of the most suitable histological procedures to study adult hippocampal neurogenesis in humans.

    To identify the most efficient sample pretreatment for eliminating background and autofluorescence caused by aldehyde fixation, we combined a range of concentrations of NaBH4 with HC-AR and 1.5% glycine (Gly) incubation protocols. ah, Representative images (ag) and number (h) of DCX+ cells detected in the different experimental conditions (one-way ANOVA, F24,96 = 6.632, P < 0.0001). The graph represents mean values ± s.e.m. Untreated sections (a) showed low signal specificity and intense nonspecific background, together with the presence of abundant autofluorescent aggregates (white arrows). These phenomena impeded the detection of DCX+ cells. We assayed a range of NaBH4 concentrations, alone or in combination with HC-AR and glycine incubation, and observed remarkable changes in signal specificity and background intensity. The best results were obtained after 30 min of incubation with 0.5% NaBH4 together with HC-AR (b,h). These conditions rendered the highest specificity of the signal, allowed observation of the morphological characteristics of the cells, and provided significantly reduced background. Similarly good results were obtained when combining 0.5% and 1% NaBH4 + HC-AR + glycine (h). However, glycine incubation was not used in subsequent experiments given that this step required an additional five washes, which increased the risk of tissue damage. In ce, increasing the concentration of NaBH4 over 2% caused an increase in background, thus preventing correct identification of DCX+ cells. Application of HC-AR alone did not reduce the background or the intensity of autofluorescent aggregates, but it did allow better detection of DCX+ cells, as a result of a subtle increase in signal intensity (f,g). i,j, Representative images of the CA1 hippocampal area showing the presence of intraneuronal autofluorescent aggregates in the green channel. The CA1 area was selected to show these aggregates because they are much more abundant in this area than in the DG. Incubation with Autofluorescence Eliminator reagent (Methods) after completion of immunohistochemistry caused a significant decrease in the signal for these aggregates, without affecting antibody detection. In aj, n = 6 control subjects. Twenty measurements were performed for each subject. Yellow scale bars, 50 μm; white scale bars, 20 μm. Green triangles indicate DCX+ cells; white arrows indicate nonspecific autofluorescent aggregates. *0.05 > P ≥ 0.01; **0.01 > P ≥ 0.001; ***P < 0.001. Asterisks indicate significant differences with respect to the samples that were not subjected to NaBH4, HC-AR, or glycine pretreatment in Tukey post hoc comparisons.

  4. Extended Data Fig. 4 Validation of DCX+ signal specificity.

    a, Four distinct anti-DCX antibodies were used. The company, host species, RRID, immunogen, concentration of use, and signal enhancement protocol applied are indicated. b, Schematic diagram of the DCX protein. The anti-DCX antibodies used were raised against different domains of the protein. c, Number of DCX+ cells detected by using the distinct anti-DCX antibodies, with or without NaBH4 and HC-AR sample pretreatment (one-way ANOVA, F8,32 = 15.07, P < 0.0001). The graph represents mean values ± s.e.m. NaBH4 + HC-AR incubation was required to adequately detect DCX+ cells for all the antibodies tested, although polyclonal goat anti-DCX (Santa Cruz) and mouse anti-DCX (Santa Cruz) antibodies gave the best results. d, Percentage of DCX+ cells double labeled by a polyclonal goat anti-DCX antibody (Santa Cruz) and other anti-DCX antibodies (one-way ANOVA, F6,24 = 27.05, P < 0.0001). The graph represents mean values ± s.e.m. e, Percentage of DCX+ cells double labeled by a monoclonal mouse anti-DCX antibody (Santa Cruz) and other antibodies (one-way ANOVA, F6,24 = 9.754, P < 0.0001). The graph represents mean values ± s.e.m. f,g, Representative images showing double labeling of DCX+ cells by using a polyclonal goat anti-DCX antibody (Santa Cruz) and a monoclonal mouse anti-DCX antibody (Santa Cruz) (f) or a polyclonal goat anti-DCX antibody (Santa Cruz) and a polyclonal rabbit anti-DCX antibody (Atlas-Sigma) (g). On the basis of signal specificity, signal/background ratio, tissue penetration, and working concentration, we opted to use the polyclonal goat anti-DCX antibody (Santa Cruz) to perform all subsequent quantifications. h, Representative image of a DCX+ cell counterstained with Nissl and visualized by immunohistochemistry. i, Dot-blot experiment showing no signal after preadsorption of polyclonal goat anti-DCX antibody (Santa Cruz) with a control synthetic DCX peptide that comprised the C-terminal domain of the protein. The full-length unprocessed blot is shown as source data. j, Representative image of DCX staining after sample incubation with preadsorbed polyclonal goat anti-DCX antibody (in red) showing no signal in the red channel. In ch, n = 5 control subjects. Twenty measurements were performed for each subject. In i and j, the experiment was repeated twice with identical results. Yellow scale bars, 50 μm; blue scale bars, 10 μm; black scale bars, 20 μm. Magenta triangles indicate DCX+ cells. *0.05 > P ≥ 0.01; **0.01 > P ≥ 0.001; ***P < 0.001. In ce, asterisks indicate changes with respect to the samples that did not receive NaBH4 + HC-AR treatment in Tukey post hoc comparisons. Source Data

  5. Extended Data Fig. 5 Expression patterns of various cell markers in the human dentate gyrus.

    aj, Representative images of the expression patterns of various cell markers in the DG, in samples with or without pretreatment with 0.5% NaBH4 and HC-AR: GFAP (a), PH3 (b), CR (c), PSA-NCAM (d), Prox1 (e), NeuN (f), 4R-tau (g), total tau (h), βIII-tubulin (i), and CB (j). Red scale bars, 50 μm. n = 13 control subjects. Ten stacks of images were analyzed for each subject. Blue color indicates DAPI staining.

  6. Extended Data Fig. 6 Density of cells expressing PSA-NCAM and DCX in the entorhinal cortex and the hippocampus of neurologically healthy subjects.

    a, Density of PSA-NCAM+ and PSA-NCAM+DCX+ cells in the EC. b, Density of PSA-NCAM+ and PSA-NCAM+DCX+ cells in the CA1 hippocampal subfield. c, Density of PSA-NCAM+ and PSA-NCAM+DCX+ cells in the DG. d, Density of PSA-NCAM+ and PSA-NCAM+DCX+ cells in the CA3 hippocampal subfield. No PSA-NCAM+DCX+ cells were found in non-neurogenic regions. In ad, n = 13 control subjects. Graphs represent mean values. e, Correlation between age of the subject and the number of PSA-NCAM+ cells in the EC (two-sided Pearson’s correlation, r = 0.0346, P = 0.911). f, Correlation between age of the subject and the number of PSA-NCAM+ cells in the CA1 hippocampal subfield (two-sided Pearson’s correlation, r = 0.0444, P = 0.885). g, Correlation between age of the subject and the number of PSA-NCAM+ cells in the DG (two-sided Pearson’s correlation, r = 0.2965, P = 0.325). h, Correlation between age of the subject and the number of PSA-NCAM+ cells in the CA3 hippocampal subfield (two-sided Pearson’s correlation, r = −0.299, P = 0.321). i, Correlation between PMD and the number of PSA-NCAM+ cells in the EC (two-sided Pearson’s correlation, r = −0.0422, P = 0.151). j, Correlation between PMD and the number of PSA-NCAM+ cells in the CA1 hippocampal subfield (two-sided Pearson’s correlation, r = −0.0916, P = 0.766). k, Correlation between PMD and the number of PSA-NCAM+ cells in the DG (two-sided Pearson’s correlation, r = −0.0584, P = 0.849). l, Correlation between PMD and the number of PSA-NCAM+ cells in the CA3 hippocampal subfield (two-sided Pearson’s correlation, r = −0.276, P = 0.360). In el, n = 13 control subjects. m, Influence of sex on the number of PSA-NCAM+ cells detected in the EC (Mann–Whitney test, U2,13 = 12.00, P = 0.4140). n, Influence of sex on the number of PSA-NCAM+ cells detected in the CA1 hippocampal subfield (Mann–Whitney test, U2,13 = 13.00, P = 0.4751). o, Influence of sex on the number of PSA-NCAM+ cells detected in the DG (Mann–Whitney test, U2,13 = 12.00, P = 0.4140). p, Influence of sex on the number of PSA-NCAM+ cells detected in the CA3 hippocampal subfield (Mann–Whitney test, U2,13 = 17.50, P = 1.000). In mp, n = 13 control subjects. Graphs represent mean values ± s.e.m.

  7. Extended Data Fig. 7 Patients with Alzheimer’s disease included in this study.

    a, Epidemiological aspects of the patients with AD included in this study. Age, sex, PMD, Braak and CERAD stage, type of fixative, and fixation time are indicated. All samples were fixed for 24 h in 4% PFA at 4 °C. Moreover, the case indicated with an exclamation mark was included in control fixation experiments (c,d), and additional blocks were thus fixed for 1, 2, 6, 12, and 48 h in the same fixative at 4 °C. b, Age distribution in the groups of subjects (Kruskal–Wallis test, K7,58 = 19.09, P = 0.0040). The graph represents mean values ± s.e.m. n = 13 control subjects and 45 patients with AD. c, Density of DCX+ cells detected in a fixation time experiment performed on samples from a patient with Braak stage VI AD (case AD 45; see a). d, Density of PSA-NCAM+ cells detected in the same patient. In c and d, cell counts were performed independently on 20 sections from case AD 45 for each fixation time point. Graphs represent mean values. The same effects as those observed in control subjects take place, namely, a reduction in staining as fixation time increases and recovery of signal after NaBH4 and HC-AR treatment. *0.05 > P ≥ 0.01; **0.01 > P ≥ 0.001; ***P < 0.001. In b, asterisks indicate significant differences with respect to control subjects in a Dunn’s multiple-comparison test.

  8. Extended Data Fig. 8 Histological alterations observed in the dentate gyrus of patients with Alzheimer’s disease.

    ag, Representative images of the DG of control subjects and patients with AD showing β-amyloid and phosphorylated tau (Ser396) staining in the DG along the six Braak stages of the disease1,20. The presence of these structures in the DG progressively increases as the disease advances. In ag, n = 13 control subjects and 45 patients with AD. h, Density of NeuN+ neurons in the GCL. No significant differences were observed during progression of the disease (one-way ANOVA test, F7,58 = 1.135, P = 0.3565). The graph represents mean values ± s.e.m. i, Ratio between the density of DCX+ cells and the number of NeuN+ neurons in the GCL (one-way ANOVA test, F7,58 = 11.08, P < 0.0001). The graph represents mean values ± s.e.m. j, Correlation between the number of DCX+ cells and the fluorescence intensity of phosphorylated tau (Ser396) in the DG (Pearson’s correlation, r = −0.5316, P < 0.0001). k, Correlation between the number of DCX+ cells and the fluorescence intensity of β-amyloid in the DG (Pearson’s correlation, r = −0.3934, P = 0.0025). In hk, n = 13 control subjects and 45 patients with AD. Twenty measurements were performed for each subject to determine either the density of DCX+ cells or the number of NeuN+ neurons. Ten measurements were performed for each subject to determine the fluorescence intensity of phosphorylated tau (Ser396) and β-amyloid. **0.01 > P ≥ 0.001; ***P < 0.001. In h and i, asterisks indicate significant differences with respect to control subjects in Tukey post hoc comparisons. In j and k, asterisks represent statistical significance in Pearson’s correlation. White scale bars, 50 μm. Blue triangles indicate β-amyloid senile plaques; magenta triangles indicate neurons that show tau phosphorylation, neurofibrillary tangles, or dystrophic neurites.

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    Full-length unprocessed Dot Blot membrane

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https://doi.org/10.1038/s41591-019-0375-9

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